FORWARD GENETICS TO UNRAVEL NOVEL GENES AND THEIR FUNCTIONS IN
TOMATO FRUIT DEVELOPMENT
By
Satya Swathi Nadakuduti

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
Plant Breeding, Genetics and Biotechnology - Horticulture - Doctor of Philosophy
2014

ABSTRACT
FORWARD GENETICS TO UNRAVEL NOVEL GENES AND THEIR FUNCTIONS IN
TOMATO FRUIT DEVELOPMENT
By
Satya Swathi Nadakuduti
Tomato (Solanum lycopersicum) is a member of the large and phenotypically diverse
Solanaceae family that includes several other agriculturally valuable crops such as potato,
eggplant, pepper, petunia and tobacco. Along with its role as an important vegetable crop, tomato
also serves as an excellent model system for studying fleshy fruit development. This dissertation
focuses on the characterization of four tomato mutants: sticky peel (pe), uniform gray-green (ug),
peach (p), and easy peel (ep) that each display altered phenotypes associated with fruit
development. The main research objectives were (1) to evaluate the phenotypes of these mutants
(2) position each locus on the tomato genetic map and isolate the mutant genes by positional
cloning and (3) functionally characterize the genes to determine their role in tomato fruit
development.
Fruit of the pe mutant have sticky and glossy fruit surface and are cutin deficient with an
altered wax profile. In addition, several phenotypes attributed to altered epidermal cell function
are apparent in pe, including altered leaf cuticle composition, leading to increased transpiration, a
lower trichome density and reduced anthocyanin accumulation. Genetic mapping revealed that
pe encodes a new allele of CUTIN DEFICIENT 2 (CD2), a member of the Class IV
homeodomain-leucine zipper transcription factor (TF) family that was previously only associated
with a fruit-specific cuticle deficiency. The Arabidopsis anthocyaninless 2 mutant encodes a
homolog of CD2 and additional characterization of this mutant revealed that it also exhibits a
mildly cutin deficient phenotype, thus revealing a regulatory link between cuticle and flavonoid

biosynthesis. This study identified a null allele of CD2 and further extended the role of CD2 to
multiple pathways operating within the epidermis that have evolved as adaptations to stress thus
revealing its role as a master regulator of epidermal cell function.
In tomato, a latitudinal gradient of chloroplast development exists, which leads to the
formation of a green shoulder at the peduncle end of the fruit that positively impacts the nutrient
content of ripe fruit. Fruit of the ug mutant lack the green shoulder surrounding the calyx.
Positional cloning revealed that UG encodes TKN4, a class I Knotted1-like homeobox (KNOX)
TF. KNOX genes typically influence plant morphology through maintenance of meristem
activity. However, this research identified a previously undefined role for these TFs in the
establishment of chloroplast gradients in tomato fruit through regulating the expression of the
GOLDEN 2-LIKE TF, SlGLK2, and it’s distant homolog ARABIDOPSIS PSEUDORESPONSE
REGULATOR-LIKE (SlAPRR2-LIKE). This study has identified an atypical role for KNOX in
regulating fleshy fruit development and defined functional interactions between well-studied, yet
previously disparate, TFs that influence fruit chloroplast development.
Fruit of the p mutant are dull, covered by dense trichomes with altered epidermal cell
structure and cuticular composition. Fruit of the ep mutant can be peeled easily with limited
damage to the underlying pericarp cells. Inter-specific crosses of p and ep mutants with S.
pimpinellifolium were performed to generate F2 mapping populations that were screened with
genetic markers to assign the mutant loci to chromosomes 2 and 8, respectively, although the
mapping is still in progress. The aim of this research is to ultimately isolate these mutant loci by
positional cloning and perform functional analysis of each gene. Together, the work presented in
this dissertation expands the current knowledge base on the molecular factors that influence fruit
development.

This work is dedicated to my loving husband, Raghu for all his support to follow my dream.

iv

ACKNOWLEDGEMENTS
My life as a graduate student for the past five years at Michigan State University has been
the most challenging and at the same time, most rewarding. I have to step back in time to thank
people who are responsible to change the scope of my education and thereby my life by inspiring
me to pursue higher education, my father and my brother. When I went to pursue my Master’s in
Hannover, Germany, Dr. Hartmut Stützel, my mentor, has greatly solidified my passion for
science and research, which naturally led me to PhD. I feel very lucky in life to marry an
amazing person, which also gave me an opportunity to come to MSU to fulfill my dream.
First, I am extremely thankful to Dr. Cornelius Barry who accepted me as his graduate
student and gave me an opportunity to get premiere education and be part of a great research
community, here at MSU. I have the privilege of being one of his first students and have learnt a
lot from him. Along with working in science, improving writing skills, the most important thing
that I learned is time management, which he emphasized right from the beginning. Preparing a
task list has now integrated into my everyday life. He has been a great mentor and provided
tremendous support at every stage of my research along with helping me immensely with my
dissertation. He always prioritized his students, which always kept me motivated at work.
In addition to Corny, I would like to render my sincere thanks to members of my
guidance committee, Dr. John Ohlrogge, Dr. Rob Last and Dr. Robin Buell for readily accepting
to be on my committee. Collaboration with John has been a great boost to my work. John has
given me great moral support and strength. Few minutes of our conversation have always
provided me much comfort and encouragement. Rob and Robin, I believe, contribute to all
graduate students by offering amazing courses that form a strong base for every student. My first
year of taking their courses has made a great impression on me. I greatly appreciate their time to
v

discuss my research updates as well as their valuable suggestions regarding my current research
and future options.
All the past and present Barry lab members have helped me along the way and created a
friendly environment in the lab. My special thanks to Bill, Chelsy and Matt for their
contributions to TKN4 and easy peel projects and Charity, for helping me in green house
especially during my pregnancy. Dr. Eliana Gonzales-Vigil has been a great friend and advisor. I
also thank Krystle, Julia, Dr. Qian Ma who always lent a helping hand. My hearty thanks to Dr.
Dylan Kosma and Dr. Mike Pollard for introducing me to the cuticle world and educating me
further. I would also like to thank John’s other lab members who were always helpful and
friendly.
More broadly, I would like to thank the Plant Breeding, Genetics, and Biotechnology
program and the Department of Horticulture. I thank all the members of joint lab for devoting
their time to listen to my research updates and for their helpful comments. I would also like to
thank the friendly staff in this department for their help. My thanks to all the friends I have made
in the university who contributed one way or the other during my time at MSU.
I feel extremely blessed to have a very closely knit family and would like to express great
love to all. Especially, my husband, Raghu who did everything he possibly could, to make my
dream of studying at MSU come true. My one year old son, Pranav brought so much joy and
fulfillment in our lives and brought the family even closer. Two amazing ladies in my life, my
mom and my mother-in-law, took great care of our baby and us during the past one year, without
whom I couldn’t have reached this stage.
Finally I would like to thank all the women in science, who are my constant inspiration.

vi

TABLE OF CONTENTS

LIST OF TABLES ........................................................................................................................ x
LIST OF FIGURES ..................................................................................................................... xi
CHAPTER I .................................................................................................................................. 1
Tomato (Solanum lycopersicum), a model crop species for studying fruit development ....... 1
Tomato genetics and genomics-based resources ................................................................... 2
Tomato fruit anatomy ............................................................................................................ 4
Genetic regulation of tomato fruit development .................................................................... 7
Forward genetics-based approaches provide insight into genes involved in fruit
development and ripening ...................................................................................................... 9
Fruit chloroplast development and its role in influencing fruit quality ............................... 10
The tomato fruit cuticle: structure and function .................................................................. 12
The 10, 16-diOH fatty acid-based cutin of tomato .............................................................. 13
Composition and functions of tomato cuticular waxes ........................................................ 18
Regulators of cuticle biosynthesis ....................................................................................... 20
REFERENCES .................................................................................................................... 23
CHAPTER II ............................................................................................................................... 38
Pleiotropic phenotypes of the sticky peel (pe) mutant provide new insight into the role of
CUTIN DEFICIENT2 in epidermal cell function in tomato ................................................... 38
Abstract ................................................................................................................................ 39
Introduction .......................................................................................................................... 40
Results .................................................................................................................................. 43
Introgression of the pe allele into the Ailsa Craig genetic background ...................... 43
Altered morphology and trichome chemistry of the pe mutant .................................. 43
Reduced anthocyanin accumulation in pe................................................................... 46
Lignin content is not altered in pe............................................................................... 47
The pe fruit cuticle has altered physical and chemical properties .............................. 47
Altered surface chemistry and increased water conductance in pe leaves .................. 52
Root suberin and wax composition are not dramatically altered in pe ....................... 57
Mapping of the pe locus and candidate gene identification ........................................ 61
Mutation at the anthocyaninless 2 locus leads to cutin deficiency ............................. 64
Discussion ............................................................................................................................ 68
Characterization of pe identifies additional roles for CD2 ......................................... 68
HD-ZIP IV proteins and cuticle biosynthesis ............................................................. 72
A role for CD2 in reducing leaf water loss ................................................................. 73
CD2 and phenylpropanoid biosynthesis ..................................................................... 75
CD2 as a regulator of epidermal cell function ............................................................ 76
Materials and Methods ......................................................................................................... 78
Plant material and growth conditions ......................................................................... 78
Extraction and analysis of cuticular lipids .................................................................. 79
Imaging of cuticular lipids and scanning electron microscopy .................................. 80
vii

Imaging anthocyanin distribution ............................................................................... 80
Determination of stomatal and trichome densities...................................................... 80
Analysis of volatile terpenes from tomato trichomes ................................................. 81
Determination of fruit glossiness ................................................................................ 81
Mechanical analysis of the fruit cuticle ...................................................................... 82
Determination of anthocyanin and chlorophyll content.............................................. 82
Determination of leaf epidermal permeability ............................................................ 82
Lignin analysis ............................................................................................................ 83
Genetic mapping ......................................................................................................... 83
Cloning of CD2 and phylogenetic analysis ................................................................ 83
Construct assembly and silencing of CD2 .................................................................. 84
qRT-PCR analysis ....................................................................................................... 84
Statistical analyses ...................................................................................................... 85
Acknowledgements .............................................................................................................. 85
REFERENCES .................................................................................................................... 89
CHAPTER III ............................................................................................................................. 98
KNOX genes influence a gradient of fruit chloroplast development through regulation of
GOLDEN2-LIKE expression in tomato ..................................................................................... 98
Abstract ................................................................................................................................ 99
Introduction ........................................................................................................................ 100
Results ................................................................................................................................ 102
Positional cloning of ug defines a role for KNOX in fruit chloroplast development 102
KNOX over-expression alters fruit development and positively impacts chlorophyll
accumulation ............................................................................................................. 108
KNOX loci establish gradients of SlGLK2 expression that impact fruit chlorophyll
accumulation ............................................................................................................. 111
The fruit phenotype of hp-1 is partially suppressed by u and ug .............................. 116
Discussion .......................................................................................................................... 118
A role for KNOX genes in fleshy fruit development................................................. 118
Differential regulation of chloroplast development in leaves and fruits of tomato .. 120
TKN4 influences the gradient of chloroplast development in tomato fruit through
regulation of SlGLK2 ................................................................................................ 121
Materials and Methods ....................................................................................................... 124
Plant material, growth conditions and chlorophyll extraction .................................. 124
Genetic mapping of the ug locus and genotyping ..................................................... 124
Molecular cloning and sequence analysis ................................................................. 125
qRT-PCR analysis ..................................................................................................... 125
Microscopy ............................................................................................................... 125
Statistical analyses .................................................................................................... 126
Acknowledgements ............................................................................................................ 126
REFERENCES .................................................................................................................. 131
CHAPTER IV............................................................................................................................ 138
Genetic mapping and morphological characterization of the tomato fruit developmental
mutants easy peel (ep) and peach (p) ....................................................................................... 138
viii

Abstract .............................................................................................................................. 139
Introduction ........................................................................................................................ 140
Results ................................................................................................................................ 143
Phenotypic characterization of the peach mutant ..................................................... 143
The genetic background of the peach mutant ........................................................... 145
Fruit of the peach mutant have altered cuticular wax composition .......................... 148
Genetic mapping of the peach locus ......................................................................... 153
Phenotypic characterization of the easy peel mutant ................................................ 156
Genetic mapping of the easy peel locus .................................................................... 156
Discussion .......................................................................................................................... 161
A range of altered fruit surface properties contributes to the p mutant phenotype... 161
An anatomical perspective of the easy peel phenotype ............................................ 162
Advancing genetic mapping strategies to identify the genes associated with ep and p
loci............................................................................................................................. 163
Materials and Methods ....................................................................................................... 166
Plant material and growth conditions ....................................................................... 166
Genetic Mapping ....................................................................................................... 166
Extraction and Analysis of Cuticular Lipids ............................................................. 167
Scanning electron microscopy .................................................................................. 169
Statistical analyses .................................................................................................... 169
REFERENCES .................................................................................................................. 170
CHAPTER V ............................................................................................................................. 176
Conclusions and perspectives................................................................................................... 176
CD2, a master regulator of epidermal cell function ........................................................... 177
Uncovering the genetic basis of fruit chlorophyll gradients .............................................. 179
Identification of loci underlying ep and p mutants will contribute to better understanding of
fruit exocarp ....................................................................................................................... 183
Importance of gene discovery in fruit crops and its impact on science and agriculture .... 185
REFERENCES .................................................................................................................. 188

ix

LIST OF TABLES

Table 2.1 Anthocyanin and chlorophyll content of AC and pe leaves ......................................... 46
Table 2.2. Lignin composition of leaves and stems in AC and pe (μg mg-1 AIR) ....................... 47
Table 2.3. Cutin monomer composition in fruit cuticles of AC and pe in μg cm-2 (%) ............... 50
Table 2.4.Total wax load in fruits of AC and pe at green and ripe stages in µg cm-2 .................. 50
Table 2.5. Fruit wax composition of AC and pe in μg cm-2 x 102 ................................................ 51
Table 2.6. PCR based genetic markers flanking the pe locus ....................................................... 86
Table 2.7. Oligonucleotide primers used in the study .................................................................. 87
Table 2.8. Homologs of CD2 utilized in phylogenetic analysis ................................................... 88
Table 3.1. Tomato accessions used in the study ......................................................................... 127
Table 3.2. PCR based genetic markers flanking the ug locus ..................................................... 128
Table 3.3. Oligonucleotide primers used in the study ................................................................ 129
Table 3.4. KNOX genes used for amino acid alignments........................................................... 130
Table 4.1. Fruit wax composition of AC and p in μg dm-2 ......................................................... 150
Table 4.2. Cutin monomer composition of WT and p in μg cm-2 (%) ........................................ 152
Table 4.3. PCR based genetic markers used for mapping p locus .............................................. 155
Table 4.4. PCR based genetic markers used for mapping ep locus ............................................ 160

x

LIST OF FIGURES

Figure 1.1. Tomato fruit anatomy ................................................................................................... 6
Figure 1.2. The structures of typical cutin monomers found in tomato fruit cutin ....................... 16
Figure 1.3. Biosynthesis of C16 family of fatty acyl monomers .................................................. 17
Figure 1.4. Tomato cuticular wax components ............................................................................. 19
Figure 2.1. Phenotypic variation between AC and pe .................................................................. 44
Figure 2.2. Comparison of relative monoterpene and sesquiterpene levels in AC and pe
trichomes ....................................................................................................................................... 45
Figure 2.3. Fruit phenotypes of pe ................................................................................................ 48
Figure 2.4. Relative fruit wax composition in AC and pe ............................................................ 53
Figure 2.5. Leaf cutin and wax profiles of AC and pe.................................................................. 55
Figure 2.6. Increased cuticular permeability in pe ........................................................................ 58
Figure 2.7. Suberin and root wax composition in AC and pe ....................................................... 59
Figure 2.8. Characterization of the pe locus and silencing of CD2 .............................................. 62
Figure 2.9. Phylogenetic and expression analysis of CD2............................................................ 65
Figure 2.10. Cutin monomer and wax profile of Arabidopsis Col-0 and anl2 leaves .................. 67
Figure 2.11. Comparison of leaf cutin and wax composition in M82 and cd2. ............................ 69
Figure 2.12. Trichome density on leaves and anthocyanin accumulation in leaves and stem peels
....................................................................................................................................................... 71
Figure 3.1. Fruit phenotype and chloroplast development in ug ................................................ 103
Figure 3.2. Genetic mapping of the ug locus .............................................................................. 105
Figure 3.3. Genotyping of TKN4 and SlGLK2 in various tomato accessions ............................. 106
Figure 3.4. Leaf phenotypes and relative expression of TKN4 ................................................... 107
Figure 3.5. Enhanced chloroplast development in Cu fruit ........................................................ 109
xi

Figure 3.6. Chlorophyll and relative expression of genes in Cu leaf .......................................... 110
Figure 3.7. Mutations at knox loci disrupt the gradient of SlGLK2 expression to impact fruit
chloroplast development ............................................................................................................. 112
Figure 3.8. Relative expression gradient of TKN2 within the fruit ............................................. 113
Figure 3.9. Silencing of SlARF4 impacts fruit chlorophyll and expression of SlGLK2 and
SlAPRR2-LIKE ............................................................................................................................ 115
Figure 3.10. The u and ug loci partially suppress hp-1 ............................................................... 117
Figure 3.11. A proposed model for KNOX activity in developing tomato fruit ........................ 122
Figure 4.1. Fruit phenotype of peach (p) LA2357 ...................................................................... 144
Figure 4.2. peach (p) fruit has altered fruit surface .................................................................... 146
Figure 4.3. Variations in peach (p) ripe fruit surface ................................................................. 147
Figure 4.4. Cuticular wax constituents of fruits of WT and p .................................................... 149
Figure 4.5. Genetic linkage map of chromosome 2 .................................................................... 154
Figure 4.6. Fruit phenotypes of easy peel (ep)............................................................................ 157
Figure 4.7. Association of C2_At5g27390 marker to ep phenotype .......................................... 158
Figure 4.8. Genetic linkage map of chromosome 8 showing the region of interest ................... 159

xii

CHAPTER I

Tomato (Solanum lycopersicum), a model crop species for studying fruit development

1

Tomato genetics and genomics-based resources
Tomato (Solanum lycopersicum) is a member of the large and phenotypically diverse
Solanaceae family that includes several other agriculturally valuable crops such as potato,
eggplant, pepper, petunia and tobacco. Tomato and its wild relatives originated in South America
(Ecuador, Peru and Chile) and is believed to have been domesticated in Central America (Rick,
1991; Peralta and Spooner, 2007). The genetic basis of domestication and artificial selection
evaluated by comparative transcriptomics of cultivated tomato and its wild relatives revealed
only very few changes at both the sequence and transcriptome level associated with this event
(Bai and Lindhout, 2007; Paran and van der Knaap, 2007; Chakrabarti et al., 2013; Koenig et al.,
2013; Bergougnoux, 2014). Tomato is a fleshy fruit bearing species and an important commodity
for both the fresh market and processing industries. In the United States, the production was 12.5
million tons in 2011 with a gross production value of $11.7 billion. With worldwide production
reaching almost 162 million tons in 2012, tomato is the seventh most important crop species after
maize, rice, wheat, potatoes, soybeans and cassava (FAOSTAT, http://faostat3.fao.org)
(Bergougnoux, 2014). Along with its role as an important crop, tomato also serves as an
excellent model system for studying a range of biological processes, including fleshy fruit
development and ripening, plant specialized metabolism, plant pathogen and pest interactions,
wound signaling etc., Due to its importance as a model crop species numerous resources have
been developed for tomato and its close relatives to facilitate research and crop improvement
(Kimura and Sinha, 2008; Menda et al., 2013).
The C.M Rick Tomato Genetics Resource Center (TGRC) in University of California,
Davis maintains seed stocks of germplasm resources including cultivars, wild relatives,
monogenic mutants, and miscellaneous genetic stocks of tomato (http://tgrc.ucdavis.edu/). Wild

2

relatives constitute a reservoir of genetic and phenotypic diversity and yield viable crosses with
cultivated tomato species. Thousands of spontaneous monogenic mutants were catalogued and
are publicly available. A comprehensive mutant library was also developed in the background of
the inbred variety M82 derived from EMS (ethyl methane sulfonate) and fast-neutron
mutagenesis which contains several thousand mutants accessible at “The Genes That Make
Tomatoes” (http://zamir.sgn.cornell.edu/mutants) (Menda et al., 2004). Recombinant inbred line
populations (Paterson et al., 1991; Goldman et al., 1995; Ashrafi et al., 2009) as well as
introgression line populations developed from wild relatives in S. lycopersicum backgrounds
(Eshed and Zamir, 1995; Monforte and Tanksley, 2000) have greatly facilitated the quantitative
trait loci (QTL) based studies. Micro-Tom, a cultivar with small stature and short generation time
has been adopted as a model system for tomato functional genomics and accelerated
transformation protocols and several genomic resources have been developed, including EMS
mutants and TILLING populations (Meissner et al., 1997; Emmanuel and Levy, 2002;
Matsukura et al., 2008).
The first “classical” linkage map of tomato, created with morphological and
physiological markers on all 12 linkage groups was developed (Butler, 1968) and was later
expanded (Rick, 1975). This facilitated the mapping of isozymes, which led to the first isozyme
linkage map (Tanksley and Rick, 1980). Subsequently, high-density molecular linkage maps
were constructed by mapping restriction fragment length polymorphism (RFLP) and amplified
fragment length polymorphism (AFLP) markers to the tomato genome (Paterson et al., 1988;
Tanksley et al., 1992; Haanstra et al., 1999). Several RFLP markers were converted into PCRbased Cleaved Amplified Polymorphic Sequence markers (Frary et al., 2005). For construction
of physical maps, Yeast artificial chromosome (YAC) libraries (Martin et al., 1992) and bacterial

3

artificial chromosome libraries (BACs) (Budiman et al., 2000) were constructed from tomato
which were successfully used for several positional cloning experiments. Tomato expressed
sequence tag (EST) clones and microarrays are available through (http://ted.bti.cornell.edu/).
Furthermore, with the advent of high throughput genotyping platforms, high density single
nucleotide polymorphisms (SNPs) arrays and high density genetic maps are now available
(Hamilton et al., 2012; Sim et al., 2012). A high quality reference genome sequence of the
‘Heinz 1706’ cultivar is now available (The Tomato Genome Consortium, 2012) and

re-

sequencing efforts for 150 tomato genomes including landraces, wild species as well as modern
varieties is ongoing and will further aid in understanding the basis of adaptation, evolution and
address the intra- and inter-specific relationships (http://www.tomatogenome.net/). Repositories
for tomato information include The SOL Genomics Network (http://solgenomics.net/),
Solanaceae Genomics Resource (http://solanaceae.plantbiology.msu.edu), the

Tomato

Functional Genomics Database (http://ted.bti.cornell.edu/) and a comprehensive list of other
genomic databases available for tomato is given in (Ronning, 2013).
Tomato fruit anatomy
Botanically, tomato fruit is a berry, which develops from a fertilized ovary and contains
seeds within a fleshy pericarp. Tomato fruit pericarp is formed from the outer ovary wall, which
forms the bulk of the fruit. The pericarp surrounds the placenta and locules. In immature green
fruit, the placental tissue is firm and fills the locules but by the mature green stage, locular tissue
becomes jelly like and contains seeds. The pericarp consists of an outer exocarp constituting the
fruit skin, parenchymatous mesocarp with vasculature and an inner endocarp. Basic anatomy of
the tomato fruit is shown in Figure 1.1. The pericarp wall may also be divided into outer walls
called septa, which separate the adjacent locules and the inner wall known as columella. The
4

number of locules is dependent on the number of carpels present in the gynoecium and can range
from two to many (Lippman and Tanksley, 2001; Munos et al., 2011). The exocarp consists of
the single cell epidermal layer that is covered by an extracellular cuticle and 3-5 layers of
hypodermal collenchyma cells. The mesocarp constitutes the largest portion of the pericarp and
consists of several layers of large vacuolated parenchyma cells and the vascular tissue.
Collenchyma and parenchyma are the main metabolic factories of fruit pericarp as they regulate
energy metabolism in the fruit. Collenchyma cells are a source of energy as they contain large
number of chloroplasts and parenchyma cells are energy sinks as they accumulate starch
granules (Matas et al., 2011). Endocarp limits the pericarp and consists of a single layer of inner
epidermal cells. There is also a cuticle covering the surface of inner epidermis that faces the
locular cavity. Unlike the cuticle covering the outer epidermis, the cuticle covering the inner
epidermis does not undergo thickening during fruit development (Matas et al., 2011). Using laser
capture micro-dissection (Matas et al., 2011) collected five principal tissues of tomato pericarp:
outer and inner epidermal layers, collenchyma, parenchyma, and vascular tissues at their
maximal growth phase and characterized the cell-specific transcriptome by pyrosequencing. This
precise method of sample collection facilitated the interrogation of the cell-specific
transcriptome profiles in developing fruit, revealing preferential expression of classes of genes in
particular cell types. For example, all the starch synthesis-related transcripts were enriched in all
fruit tissues except inner epidermis, suggesting that there could be photosynthetic capacity in all
tissues. However, transcript abundance of genes specifically related to photosynthesis,
photorespiration and photosystems are enriched in collenchyma cells, present just beneath the
outer epidermis but not in interior layers including parenchyma cells suggesting that
photosynthesis

might

be

restricted

to

outer

5

collenchyma

(Matas

et

al.,

2011).

Figure 1.1. Tomato fruit anatomy A) Cross section of mature green (MG) tomato fruit. The red
box across the pericarp represents the section shown in B) Free hand section of MG tomato fruit
stained with toluidine blue showing different layers of fruit pericarp along with vascular bundles
(VB). The yellow box represents exocarp region, which is further expanded in C) The Exocarp
and mesocarp regions are shown specifying their cell types. D) Scanning electron micrograph
showing outer epidermis of the tomato fruit with thick fruit cuticle. (Scale bar for B = 2mm; C =
100μm and D = 20μm).

6

Transcripts associated with cuticle metabolism were specific to the outer epidermis, although
some were also expressed in the inner epidermis.
Genetic regulation of tomato fruit development
Fruits are essential component of human diet and are specialized organs unique to plants.
Tomato fruit development can be divided into four stages from floral initiation to fruit ripening.
Phase I is a 2-3 week period where floral initiation and production of mature flower occurs. It is
during this phase that the identity, number as well as shape of floral organs are determined. This
phase requires the coordinated spatial and temporal expression of transcription factors, including
MADS-box genes, originally characterized in Arabidopsis and Antirrhinum majus that specify
floral organ identity and gave rise to the ABC model of floral identity (Bowman et al., 1989;
Schwarz-Sommer et al., 1990; Coen and Meyerowitz, 1991; Gustafson-Brown et al., 1994;
Ferrandiz et al., 2000; Pelaz et al., 2000; Pinyopich et al., 2003; Bowman et al., 2012). Phase II
is an active cell division period in the ovary beginning at anthesis and continues for the first 7-10
days after successful fertilization. This phase is characterized by the interplay of hormonal
signals, primarily involving auxin and gibberellins that promote the expression of genes involved
in cell division and expansion (Kibler and Bangerth, 1982; Gillaspy et al., 1993; de Jong et al.,
2009; Seymour et al., 2013). Phase III is a period of 6-7 weeks (end of cell division until a week
before onset of ripening) where fruit growth occurs through cell expansion. This phase results in
a remarkable increase in the size of the fleshy fruit. Phase IV is the ripening phase that initiates
after growth has stopped and the seed has matured. This phase involves rapid chemical and
structural changes that determine fruit aroma, color, texture, and biochemical composition (e.g.,
acids and sugars) but not fruit size and shape and is accompanied by rapid changes in
transcriptional reprogramming (Gillaspy et al., 1993; Alba et al., 2005). Interestingly, many of
7

the regulators of fruit ripening are shared with genes involved in fruit development and
dehiscence in Arabidopsis, suggesting evolutionary conserved mechanisms regulate these
processes. MADS box family transcription factors (TFs) serve as central regulators of fruit
development and ripening, not only in Arabidopsis and tomato but also in several other fruit
species (Boss et al., 2002; Tadiello et al., 2009; Klee and Giovannoni, 2011; Seymour et al.,
2011; Handa et al., 2012). SHATTERPROOF (SHP) SHP1, 2 in Arabidopsis specify valve
margin formation and dehiscence for controlled seed dispersal (Ferrandiz et al., 2000; Liljegren
et al., 2000) and its tomato ortholog, TAGL1 (AGAMOUS-LIKE1) is involved in fleshy fruit
expansion and ripening (Vrebalov et al., 2009). Similarly, APETALA2 in Arabidopsis affects
silique development by negatively regulating replum formation and valve margin lignification
(Ripoll et al., 2011), and its tomato ortholog, SlAP2a is a negative regulator of fruit ripening
(Karlova et al., 2011; Chung et al., 2012). RIPENING INHIBITOR is a master regulator of
ripening (Vrebalov et al., 2002), and is a member of the SEPALLATA sub-family of MADS-box
genes that are involved in growth and development of floral organs including sepals (Pelaz et al.,
2000; Zahn et al., 2005). FRUITFULL (FUL) in Arabidopsis plays a role in silique development
by promoting cell expansion in valves causing elongation of the silique (Gu et al., 1998). Tomato
contains two close homologs of FUL, FUL1 and FUL2, which are functionally redundant and
down regulation of both impairs color development in the fruit and affects fruit ripening
processes (Bemer et al., 2012; Shima et al., 2013). Thus, members of this family of TFs share
functional similarities.
When an ovary develops into a fruit, the ovary wall becomes the pericarp. The fruit
pericarp is derived from cells within interna1 layer L3 in the shoot apical meristem (SAM). The
number of cells that make up L3 determine the size of the floral meristem and carpel number,

8

which in turn governs the size of the pericarp. This is demonstrated from studies with interspecific chimeras between S. lycopersicum and S. peruvianum genotypes differing in the number
of carpels per flower and also chimeras generated by grafting using periclinal chimeras differing
in their genetic makeup in meristem cell layers. Thereby, as pericarp forms the bulk of the fruit,
L3 can directly determine the eventual sink size and strength of the developing fruit whereas L1
and L2 contribute to the outer and inner epidermal cell layers (Szymkowiak and Sussex, 1992;
Huala and Sussex, 1993).
Forward genetics-based approaches provide insight into genes involved in fruit
development and ripening
Many insights into the molecular basis of fruit development and ripening have been
achieved through the characterization of monogenic mutants and exploitation of natural variation
coupled with QTL analysis. Complex traits such as yield, shape and nutritional quality are
quantitative traits influenced by multiple genes. Two of the early fruit-related QTLs identified
were related to yield. The fruit weight 2.2 locus was identified as a key factor in the evolution of
fruit size during tomato domestication and the brix 9-2-5 QTL, which influences sugar content
was localized to an invertase gene (Frary et al., 2000; Fridman et al., 2000). Subsequently,
multiple studies have mapped and identified QTLs for various fruit characteristics in several
segregating populations derived from inter-specific crosses, including those associated with fruit
size, shape, weight, soluble solid content, pH, color, and fruit firmness (Grandillo et al., 2013).
Insight into fleshy fruit development and ripening has also been achieved through the
characterization and identification of loci encoded by numerous monogenic mutants of tomato
that possess altered fruit phenotypes. These studies have identified master regulators of ripening
encoded by the ripening inhibitor and Colorless non-ripening loci (Robinson and Tomes, 1968;
Vrebalov et al., 2002; Manning et al., 2006), loci involved in ethylene response (Lanahan et al.,
9

1994; Wilkinson et al., 1995; Barry et al., 2005; Barry and Giovannoni, 2006), carotenoid and
flavonoid biosynthesis (Mustilli et al., 1999; Ronen et al., 2000; Isaacson et al., 2002; Liu et al.,
2004; Adato et al., 2009), cuticle biosynthesis (Isaacson et al., 2009; Nadakuduti et al., 2012;
Yeats et al., 2012) and chloroplast development (Barry et al., 2012; Powell et al., 2012).
Fruit chloroplast development and its role in influencing fruit quality
Fleshy fruits are generally defined as a sink tissue, although fruit photosynthesis can
contribute up to 20% of the total fruit photosynthate (Hetherington et al., 1998). Chloroplast
development and the carbohydrate status of unripe fruits impact the composition and quality of
ripe fruit (Hobson and Bedford, 1989; Powell et al., 2012; Sagar et al., 2013). Despite its
importance in defining ripe fruit quality, the genetic mechanisms that control fruit chloroplast
development are not completely understood. However, emerging evidence suggests that plastid
development is differentially regulated in fruit and leaf tissues of tomato and in fruit, is
controlled by three transcription factors (TFs), (Powell et al., 2012; Pan et al., 2013; Sagar et al.,
2013; Nguyen et al., 2014) a tomato GOLDEN 2-LIKE homolog designated, SlGLK2, a tomato
homolog of ARABIDOPSIS PSEUDO RESPONSE REGULATOR 2-LIKE (SlAPRR2-LIKE), and
AUXIN RESPONSE FACTOR 4 (SlARF4). Fruit chloroplast development is positively influenced
through the action of SlGLK2 and SlAPRR2-LIKE, whereas auxin, acting through SlARF4,
negatively impacts this process (Powell et al., 2012; Pan et al., 2013; Sagar et al., 2013).
Fruits of SlARF4 / DR12 -antisense lines are dark green when immature due to increase
in chloroplast number and enhanced chloroplast development. In addition, ARF4 / DR12 antisense fruits display an unusual pattern of cell division in the pericarp, where cells continue to
divide throughout development, unlike in wild type where this process ceases 7-10 days after
fertilization. Fruits are blotchy when ripe and display increased firmness and density, enhanced
10

sugar content and prolonged shelf-life with reduced water loss at the ripe stage (Jones et al.,
2002; Sagar et al., 2013; Sagar et al., 2013).
The uniform ripening (u) locus regulates the formation of an amorphous dark green
shoulder that surrounds the calyx end of tomato fruit. u mutants have lower chlorophyll levels in
green fruit which translates to reduced carotenoid and sugar contents in ripe fruits. U encodes
SlGLK2 that belongs to GARP family of Myb TFs (Powell et al., 2012). GLK TFs regulate
chloroplast differentiation through promotion of nuclear-encoded photosynthesis-related gene
expression (Fitter et al., 2002; Yasumura et al., 2005; Waters et al., 2008; Waters et al., 2009). In
C3 plants, pairs of GLKs often act redundantly to regulate chloroplast development, whereas in
C4 plants, including maize, each GLK has a specific role in bundle sheath and mesophyll cells
that is largely determined by their differential expression (Langdale and Kidner, 1994; Rossini et
al., 2001; Waters et al., 2008; Waters et al., 2009; Wang et al., 2013). In tomato, SlGLK2
determines both chlorophyll accumulation as well as distribution within developing fruit.
Breeders have selected for the u locus to achieve uniform ripening in fruits, which results in
lower chlorophyll content and thereby less efficient in photosynthesis and compromised
carotenoid and carbohydrate levels in the ripe fruit (Powell et al., 2012). Tomato possesses two
GLKs, SlGLK1 and SlGLK2 that are functionally equivalent, however differential expression of
these genes ensures that SlGLK2 functions in fruits while SlGLK1 functions primarily in leaf
tissue (Nguyen et al., 2014). In addition, the expression of SlGLK2 is elevated in the stem end of
tomato fruit and promotes the gradient of chloroplast development that leads to the formation of
the green-shoulder that surrounds that calyx of the fruit. The chloroplast gradient and ripening
gradients were further investigated by RNA-seq analysis of three sections made along the
vertical axis of the fruit. Together with the expected chlorophyll biosynthetic genes whose

11

expression is enriched in the stem end, additional genes, including the KNOX transcription
factors, TKN2 and TKN4 were also up-regulated in this region, suggesting that they may play a
role in either fruit chloroplast development and / or the formation of gene expression gradients in
tomato fruit (Nguyen et al., 2014).
The tomato fruit cuticle: structure and function
The cuticle is considered one of the most important adaptation strategies for the
colonization of plants to terrestrial environments owing to its ability to retain water and protect
against ultra-violet radiation (Kolattukudy, 1980; Edwards, 1993; Kolattukudy, 2001). In
addition the cuticle withstands biotic stresses, such as insect and pathogen infestations, and plays
a major role in plant morphogenesis by preventing cell-cell adhesion and providing structural
support by withstanding turgor pressure from the underlying cells (Kolattukudy, 1980; Sieber et
al., 2000; Riederer and Schreiber, 2001; Wellesen et al., 2001; Nawrath, 2006; Pollard et al.,
2008; Yeats and Rose, 2013).
In tomato, the exocarp, also referred to as the fruit skin is the outermost layer of the fruit
pericarp comprised of the cuticle, outer epidermis and 3-5 layers of sub-dermal collenchyma
cells (Bargel and Neinhuis, 2005). The mechanical performance of the exocarp, mainly attributed
to the cuticle, has physiological and economic significance due to its impermeable cuticle
covering that helps to minimize water loss, pathogen infection, reduce the incidence of cracking
and provide color and gloss to the fruit surface (Chu and Thompson, 1972; Emmons and Scott,
1997; Emmons and Scott, 1998; Matas et al., 2004; Dominguez et al., 2011). The cuticle is
synthesized by the epidermal cells and is deposited at the surface of these cells and is associated
with a network of embedded polysaccharides that extend from their cell walls. The cuticle is
heterogeneous and is comprised of a cutin matrix together with intra- and epicuticular waxes.
12

Cutin is a polymer formed by inter-esterification of hydroxy fatty acids with small amounts of
glycerol, phenylpropanoids and dicarboxylic acids (Kolattukudy, 2001). The intra-cuticular
waxes are embedded in the cutin framework while the epi-cuticular waxes are present on the
surface. The cuticle can be demarcated as a ‘cuticle layer’ that is close to the epidermis
consisting of cutin and polysaccharides and an overlying ‘cuticle proper’ consisting of primarily
cutin and waxes at the air-fruit interface (Walton, 1990; Heredia, 2003). Arabidopsis has served
as the primary model to decipher several steps in cutin and wax biosynthesis, but with the
exception of its flower cutin, Arabidopsis has an atypical cutin composition compared to other
species (Bonaventure et al., 2004; Franke et al., 2005; Li-Beisson et al., 2009). Tomato fruit
possesses a thick prominent astomatous cuticle which is primarily made of a typical cutin
monomer 10, 16-dihydroxyhexadecanoic acid that comprises greater than 80% of the cutin by
weight (Vogg et al., 2004; Kosma et al., 2010) and more recently, tomato has become
increasingly utilized to study aspects of cuticle biosynthesis and function.
The 10, 16-diOH fatty acid-based cutin of tomato
Tomato cutin is dominated by a C16 family of mostly ω-hydroxy fatty acids with mid
chain hydroxyl groups or epoxy group and dicarboxylic acids making up to 85% of the total
cutin. However, similarly modified C18 fatty acids are also detected in tomato cutin, which
together with cis and trans-coumaric acids make up the remainder of the cutin (Isaacson et al.,
2009; Kosma et al., 2010; Nadakuduti et al., 2012). The fatty acids constituting typical tomato
fruit cutin monomers are listed in Figure 1.2. The pathway leading to formation of C16 type of
cutin is fairly well understood through recent advancements made using both Arabidopsis and
tomato. De novo fatty acid synthesis occurs exclusively inside the chloroplasts of epidermal
cells, which supplies acyl chains for polymer assembly (Ohlrogge and Jaworski, 1997). Further
13

modifications to the acyl chains occur outside the chloroplasts in the endoplasmic reticulum. In
Arabidopsis, hydroxylation of C16 fatty acid at the terminal position to yield ω-hydroxy C16
fatty acids is carried out by CYP86A4 (Li-Beisson et al., 2009), a tomato homolog of this
enzyme, designated SlCYP86A69, was recently identified by cloning the cutin deficient 3 (cd3)
mutant. Mutants at the slcyp86a69 locus display reduced cuticle thickness with altered cutin
composition (Shi et al., 2013). Further hydroxylation in the mid chain position to produce the
major monomer, 10, 16-dihydroxy C16 fatty acid is carried out by CYP77A6, a mid-chain
hydroxylase (Li-Beisson et al., 2009). A transcriptome study of tomato fruit pericarp identified
two CYP77A genes that are highly expressed in the outer epidermis, suggesting that they may be
candidates for mid-chain hydroxylases (Matas et al., 2011). The steps involving the formation of
dicarboxylic acid and epoxy acids are currently unclear.
Once the modified fatty acids are produced, these cutin fatty acyl precursors are linked to
glycerol by glycerol-3-phosphate acyltransferases (GPATs). For cutin biosynthesis the unique bifunctional acyl transferases, GPAT4 and GPAT6 are required. These enzymes act as
acyltransferases to esterify acyl groups, predominantly to the sn-2 position of glycerol 3
phosphate and also possess phosphatase activity that yields sn-2 monoacylglycerol (2-MAG). It
was predicted that 2-MAG could be a potential cutin precursor that is exported outside of the
epidermal cells for external cutin assembly (Li et al., 2007; Yang et al., 2010). The ATP binding
cassette (ABC) transporter ABCG12 is implicated in the transport of cutin monomers (Pighin et
al., 2004; Bird et al., 2007). Although cutin is a polymer, the steps required for its assembly,
including the mechanism of inter-esterification of hydroxy fatty acids and glycerol, the site of
polymerization, and the molecules that are exported from epidermal cells to their exterior
surfaces remained unresolved (Pollard et al., 2008; Beisson et al., 2012). Recently, identification

14

of CD1 by map based cloning of tomato cutin deficient 1 mutant provided answers to some of
these questions. CD1 encodes a member of the glycine–aspartic acid–serine-leucine motif
lipase/hydrolase (GDSL) super family of enzymes. Immunolocalization of CD1 in fruit
demonstrated that it is exclusively localized within the cuticle, outside of the epidermal cells
while CD1 transcripts preferentially accumulate within epidermal cells. The detection of the
MAG species 2-mono (10, 16-dihydroxyhexadecanoyl) glycerol in the wax fraction of cd1 fruit,
further strengthened the existing hypothesis that 2-MAG could be a precursor for cutin synthesis
that is exported outside the epidermal cell and this mechanism is probably disrupted in cd1.
Purified recombinant CD1 protein catalyzed acyltransferase reactions in vitro, resulting in
formation of up to seven 10, 16-dihydroxyhexadecanoyl monomers whereas, site-directed
mutagenesis of the conserved catalytic serine to alanine eliminated this activity. Together, these
results indicate that CD1 is a cutin synthase which catalyzes cutin polymerization outside the cell
using a 2-MAG species as acyl donor and the growing polymer chain as an acyl acceptor (Yeats
et al., 2012). Although cd1 is a null mutant (Yeats et al., 2012), fruit are not completely cutin
deficient, indicating the presence of other cutin synthases or functionally similar proteins that are
yet to be identified (Isaacson et al., 2009). Solution state NMR analysis using in-vitro
synthesized CD1 protein indicated that it catalyzes primarily the formation of linear cutin
oligomers, thus the question of branching and cross-linking in the cutin polymer remains
unresolved (Yeats et al., 2014). The identified steps in the cutin biosynthetic pathway pertaining
to tomato cutin are presented in Figure1.3.

15

Figure 1.2. The structures of typical cutin monomers found in tomato fruit cutin. The free
fatty acid form is given for each monomer, although in vivo, these monomers may also be
present as acyl-CoAs or glycerolipids. The structures of tomato fruit cutin monomers reported in
(Nadakuduti et al., 2012) are presented here.

16

Figure 1.3. Biosynthesis of C16 family of fatty acyl monomers. The free fatty acids formed in
the chloroplast undergo modifications in the endoplasmic reticulum to produce cutin monomers.
The terminal and mid-chain hydroxylation reactions are carried out by CYP450s. A plantspecific bi-functional GPAT6 enzyme esterifies 10, 16-dihydroxyhexadecanoyl group, to sn-2
position of glycerol 3-phosphate yielding 2-mono (10, 16-dihydroxyhexadecanoyl) glycerol (2MHG), an initial intermediate precursor for cutin polymer assembly is the form in which it is
exported outside the epidermal cell. CD1 catalyzes the polymerization of these monomers by
transferring the hydroxyacyl group from 2-MHG to the growing polymer external to epidermal
cell. In-vivo, substrates may be acyl-CoAs (R = CoA), free fatty acids (R = H) or glycerolipids.
The sequential order of these reactions is inferred but not experimentally determined (Li-Beisson
et al., 2009; Beisson and Ohlrogge, 2012; Yeats and Rose, 2013).

17

Composition and functions of tomato cuticular waxes
Cuticular wax is a complex mixture of a series of very-long-chain (VLC) aliphatic lipids,
triterpenoids, and minor compounds, such as sterols and flavonoids (Kunst and Samuels, 2003).
Forward and reverse genetics approaches as well as epidermal transcriptome analysis in
Arabidopsis have led to molecular characterization of enzymes catalyzing major steps in wax
biosynthesis and export, contributing to our understanding of the importance of waxes in plants
(McNevin et al., 1993; Suh et al., 2005; Kunst and Samuels, 2009; Bernard and Joubes, 2013;
Lee and Suh, 2013). Cuticular waxes are involved in regulating water relations in plants by
forming a waterproof barrier. In plants, transpirational water loss is mainly regulated by stomates
and to a certain extent by cuticular wax. As tomato fruit lacks stomata, cuticular wax becomes
crucial for water regulation (Vogg et al., 2004). Tomato fruit wax consists of very long chain
fatty acids (C20-C32), straight chain (C23-C33) and branched alkanes (C30-C32), alkenes (C33),
alcohols (C20-32), triterpenoids, sterols and cis, trans-coumarates, although it is dominated in
abundance by very long chain alkanes and triterpenoids (Bauer et al., 2004; Kosma et al., 2010;
Nadakuduti et al., 2012). Although many components of tomato wax are known, up to 15 percent
of the total wax composition remains unknown (Nadakuduti et al., 2012). A representative from
each wax class is provided in Figure 1.4. The function of cuticular waxes in tomato fruit was
demonstrated by analyzing mutants with altered cuticle properties. Fruits of the lecer6 mutant
lack a β-ketoacyl-CoA synthase, a component of the VLC fatty acid elongase complex (Leide et
al., 2007). The lecer6 mutant fruit show reduced n-alkanes in the fruit wax, which directly
correlates with their water loss. In the ‘Delayed Fruit Deterioration’ (DFD) cultivar of tomato,

18

Figure 1.4. Tomato cuticular wax components. The structure of a representative compound
belonging to each class of tomato fruit cuticular wax is provided.

19

fruits undergo normal ripening, but remain firm with no loss of integrity for remarkably long
periods after they ripen (Saladie et al., 2007). Characterization of this cultivar revealed that,
although polysaccharide components of the cell wall are critical for fruit texture, alterations in
cuticle architecture is also an integral component of the fruit ripening process. The total amount
of wax was higher in DFD than was observed in wild type with a 5-fold increase in n-alkadienes
observed but no increase in n-alkanes. There was also a doubling of the total cutin in ripe fruits
of the DFD compared to wild type. These studies show the importance of fruit cuticular waxes in
mediating water relations in tomato fruit.
Regulators of cuticle biosynthesis
The regulation of cuticle synthesis is complex as it is influenced by various factors,
including several biotic and abiotic stresses (Aharoni et al., 2004; Chassot et al., 2007; Kosma et
al., 2009; Seo et al., 2009; Kosma et al., 2010; Seo and Park, 2010; Bourdenx et al., 2011). The
epidermal cells are highly specialized and serve as the site of synthesis for cutin precursors and
wax biosynthesis and their proximity to the adjacent environment influences the expression of
epidermal cell-specific transcription factors and thereby metabolic activity of these cells (Javelle
et al., 2011). Regulatory genes involved in cuticle biosynthesis have been identified in both
Arabidopsis and tomato. WAX INDUCER1/SHINE1 (WIN1 / SHN1), SHN2 and SHN3 form a
small, distinct group of proteins belonging to the APETALA2/ethylene-responsive factor
transcription factor family (Aharoni et al., 2004; Broun et al., 2004; Kannangara et al., 2007; Shi
et al., 2011). Over-expression of WIN1/SHN1 in Arabidopsis increases epicuticular wax
biosynthesis, which led to a characteristic shiny leaf phenotype and plants that display increased
drought tolerance (Kannangara et al., 2007). However, the phenotypes associated with
WIN1/SHN1 over-expression were not restricted to wax accumulation as an increase in cutin was
20

also observed together with altered epidermal cell structure, altered trichome number and
branching and an atypical stomatal index (Aharoni et al., 2004; Broun et al., 2004; Kannangara
et al., 2007). Recently, SlSHN3 of tomato has been characterized through over-expression and
RNAi lines (Shi et al., 2013). Silencing of SlSHN3 resulted in reduced cutin and wax content of
tomato fruit cuticles due to down regulation of an array of cutin metabolism genes. For example,
the CYP450 family members SlCYP77A1, SlCYP86A68 and SlCYP86A69, Arabidopsis
homologs of which were previously shown to catalyze fatty acid hydroxylation (Li-Beisson et
al., 2009), were down-regulated. Concomitant with altered cuticle composition, SlSHN3 RNAi
lines displayed increased water loss and decreased resistance to fungal infection. However, the
impact of SlSHN3 silencing may extend beyond cuticle biosynthesis as genes related to
epidermal patterning were also down-regulated compared to untransformed lines (Shi et al.,
2013).
In tomato, characterization of the cd2 mutant led to identification of CUTIN DEFICIENT
2, a homeodomain-leucine zipper IV transcription factor (HD-ZIP IV) involved in cuticle
biosynthesis in fruits (Isaacson et al., 2009). HD-ZIP IV TFs are master regulators of epidermal
cell function (Ingram et al., 1999; Kubo et al., 1999; Nakamura et al., 2006; Javelle et al., 2010;
Javelle et al., 2011). Fruits of the cd2 mutant are severely cutin deficient and also have an altered
wax profile. Characterization of the sticky peel (pe) mutant which carries a second mutant allele
at the cd2 locus indicates that CD2 is a master regulator of epidermal cell function in tomato,
with functions that extend beyond fruit cuticle biosynthesis. For example, leaves of the pe mutant
are also cutin deficient and possess increased water permeability. In addition, characterization of
pe revealed that CD2 also influences several additional epidermal cell-related phenotypes,

21

including trichome differentiation, anthocyanin accumulation (Nadakuduti et al., 2012) together
with altered sugar and glycoalkaloid profiles in fruit (Kimbara et al., 2012).
CD2 and SlSHN3 represent tomato TFs that have pleiotropic affects on epidermal cell
function, including cuticle biosynthesis. Other MYB TFs that regulate cuticle biosynthesis have
been identified in Arabidopsis including, MYB106, MYB16 (Oshima et al., 2013) along with
MYB30 and MYB96 that are biotic and abiotic stress responsive (Raffaele et al., 2008; Seo et
al., 2009). WXP1, an AP2-domain TF that regulates wax biosynthesis and confers drought
tolerance has been identified in Medicago sativa (Zhang et al., 2005). Homologs of all the TFs
identified in other species are yet to be characterized in tomato. The transcriptional network that
regulates cuticle biosynthesis has yet to be clearly established through investigation of the
downstream targets for each TF to reveal their function in plant development and epidermal cell
function. Although several advances have been made in cuticle biology, aspects such as the
structure of the cutin polymer, intra and extra cellular transport mechanisms of cutin monomers
and waxes, enzymes for the synthesis of dicarboxylic acids, epoxy acids, and alkanes together
with the complete polymerization of cutin, the order of the reactions and substrates specificities
of the enzymes remain to be determined (Pollard et al., 2008; Beisson et al., 2012; Yeats and
Rose, 2013).

22

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37

CHAPTER II

Pleiotropic phenotypes of the sticky peel (pe) mutant provide new insight into the role of
CUTIN DEFICIENT2 in epidermal cell function in tomato

Satya Swathi Nadakuduti1, Mike Pollard2, Dylan K. Kosma2, Charles Allen Jr.1, John B.
Ohlrogge2 and Cornelius S. Barry1*
Plant Physiology (2012): 159(3): 945-60

38

Abstract
Plant epidermal cells have evolved specialist functions associated with adaptation to
stress. These include the synthesis and deposition of specialized metabolites such as waxes and
cutin together with flavonoids and anthocyanins, which have important roles in providing a
barrier to water loss and protection against UV radiation, respectively. Characterization of the
sticky peel (pe) mutant of tomato revealed several phenotypes indicative of a defect in epidermal
cell function, including reduced anthocyanin accumulation, a lower density of glandular
trichomes and an associated reduction in trichome derived terpenes. In addition, pe mutant fruit
are glossy and peels have increased elasticity due to a severe reduction in cutin biosynthesis and
altered wax deposition. Leaves of the pe mutant are also cutin deficient and the epicuticular
waxes contain a lower proportion of long chain alkanes. Direct measurements of transpiration,
together with chlorophyll leaching assays, indicate increased cuticular permeability of pe leaves.
Genetic mapping revealed that the pe locus represents a new allele of CUTIN DEFICIENT 2
(CD2), a member of the Class IV Homeodomain-Leucine Zipper gene family, previously only
associated with cutin deficiency in tomato fruit. CD2 is preferentially expressed in epidermal
cells of tomato stems and is a homolog of Arabidopsis ANTHOCYANINLESS 2 (ANL2). Analysis
of cuticle composition in leaves of anl2 revealed that cutin accumulates to approximately 60
percent of the levels observed in wild-type Arabidopsis. Together, these data provide new insight
into the role of CD2 and ANL2 in regulating diverse metabolic pathways and in particular, those
associated with epidermal cells.

39

Introduction
Plants are continually exposed to environmental changes that impact their fitness and
survival. Changes in light intensity, light quality, temperature and water availability occur on a
daily basis and insect pests and pathogens pose a constant threat. Consequently, plants have
evolved a suite of physical and chemical adaptations and defenses against abiotic and biotic
stresses that have facilitated their colonization of diverse environments. The surface properties of
plants are crucial to their successful adaptation to stress. The epidermal cell layer and associated
structural appendages such as trichomes, together with the cuticle form the primary physical and
chemical barriers that protect plants against multiple stresses (Glover, 2000; Sieber et al., 2000;
Schilmiller et al., 2008; Javelle et al., 2011). Furthermore, the surface properties of plants are
dynamic and can change in response to stress, therefore influencing the physiology of the plant
(Lu et al., 1996; Ingram et al., 2000; Abe et al., 2003; Ingram, 2008; Kosma et al., 2009; Wang et
al., 2011).
The metabolism of epidermal cells is programmed for the synthesis of lipids that form the
cuticle, a heterogeneous lipid-based barrier comprised of cutin, intra and epicuticular waxes and
polysaccharides that covers the aerial surfaces of all terrestrial plants. In addition to serving as
the principal barrier to water loss (Kolattukudy, 1980; Riederer and Schreiber, 2001; Pollard et
al., 2008) the cuticle provides structural support, possesses anti-adhesive properties that limit
pathogen infection, resists insect feeding and oviposition, and has reflective properties that
reduce heat load and limit the effect of UV-radiation (Bargel et al., 2006). Furthermore, the
chemical properties of the cuticle are dynamic with both cuticle deposition and wax composition
altering in response to water stress and abscisic acid (ABA) (Kosma et al., 2009; Wang et al.,
2011). The epidermal and sub-epidermal cells are also sites of phenylpropanoid biosynthesis, and

40

in particular the synthesis and accumulation of flavonoids and anthocyanins (Martin and Gerats,
1993; Hichri et al., 2011; Matas et al., 2011). Anthocyanins protect plants through their UV
radiation absorbing properties and their synthesis is influenced by stress and can vary with light
intensity and wavelength (Beggs et al., 1987; Li et al., 1993; Christie et al., 1994; Landry et al.,
1995; Fuglevand et al., 1996; Steyn et al., 2002). Together, these studies highlight the
importance of the epidermal cell layer in the synthesis of compounds important for stress
adaptation in plants that likely contributed to colonization of the terrestrial environment by early
land plants (Edwards et al., 1996; Cooper-Driver, 2001).
Many enzymes, transporters and regulatory factors required for the biosynthesis and
deposition of cuticular lipids have been identified and characterized in Arabidopsis (Li et al.,
2007; Panikashvili et al., 2007; Samuels et al., 2008; Panikashvili et al., 2009; Li-Beisson et al.,
2010; Li et al., 2010; McFarlane et al., 2010; Seo et al., 2011; Wu et al., 2011). Tomato fruit also
serve as a powerful system for investigating the physical properties and biosynthesis of the
cuticle fostered in large part by the abundance of cuticular lipid deposited during fleshy fruit
development, its astomatous nature and relevance in fruit cracking and postharvest storage
(Emmons and Scott, 1997; Emmons and Scott, 1998; Matas et al., 2004; Bargel and Neinhuis,
2005). Genomics- and proteomics-based approaches have identified genes and proteins
preferentially expressed in the fruit peel, and perturbation of several genes through mutagenesis
has revealed altered fruit cuticle phenotypes (Vogg et al., 2004; Hovav et al., 2007; Mintz-Oron
et al., 2008; Isaacson et al., 2009; Yeats et al., 2010; Matas et al., 2011).
Fruit of the sticky peel (pe) mutant of tomato have a “rubbery” surface texture rather than
the typical smooth surface associated with wild-type fruit rendering fruits “sticky” to the touch
(Butler, 1952). In addition, pe fruits have a highly glossy fruit surface, a phenotype recently

41

associated with cutin deficiency (Isaacson et al., 2009). In this study, a combination of
microscopy and chemical analysis revealed that pe fruit are cutin deficient and have an altered
wax profile. In addition, several phenotypes attributed to altered epidermal cell function are
apparent in pe, including cutin deficiency and altered wax deposition in leaves, together with
increased cuticular permeability, lower trichome density and reduced anthocyanin accumulation.
Genetic mapping indicated that pe encodes a new allele of CUTIN DEFICIENT 2 (CD2), which
encodes a member of the Class IV Homeodomain-leucine zipper (HD-ZIP IV) family, and was
previously only associated with a fruit-specific reduction in cuticle biosynthesis (Isaacson et al.,
2009). In addition, we show that mutation in a CD2 homolog at the anthocyaninless 2 (anl2)
locus of Arabidopsis also causes a cutin deficient phenotype in rosette leaves. Together, these
data identify additional roles for CD2 and ANL2, defining a regulatory link between cuticle and
flavonoid biosynthesis, two pathways that operate within epidermal cells that are critical for
plant responses to stress.

42

Results
Introgression of the pe allele into the Ailsa Craig genetic background
At the initiation of this research, the genetic background of the pe mutant was unknown.
Therefore, we initiated a backcross strategy to introgress the pe mutation into the Ailsa Craig
(AC) genetic background, a cultivar previously selected for introgression of varied
morphological mutants of tomato (Darby et al., 1978). The mutant and wild-type plants used in
the following experiments were derived from a BC2F2 population that we estimate contains
approximately 93.75% of the AC genetic background. Therefore, the influence of the genetic
background on the data presented is likely to be minimal.
Altered morphology and trichome chemistry of the pe mutant
In addition to the previously documented sticky peel phenotype of pe fruit (Butler, 1952),
the pe mutant exhibits a short stature with pale green leaves and stems (Figure 2.1A). Five weekold wild-type Ailsa Craig (AC) plants had an average dry weight of 6.45 g whereas the dry
weight of pe plants was 3.86 g. Glandular trichomes are epidermal cell appendages that are
prevalent on the surface of tomato plants and synthesize a suite of specialized metabolites
known, or hypothesized to play a role in plant defense responses against pests and pathogens
(Schilmiller et al., 2008). Type VI glandular trichomes are the most abundant glandular
trichomes on the tomato leaf surface (Kang et al., 2010) and these are reduced greater than threefold in pe (Figure 2.1B-D). The type VI glandular trichomes of cultivated tomato synthesize a
mixture of mono- and sesqui-terpenes of which β-phellandrene predominates (Schilmiller et al.,
2009). The volatile terpene levels in leaf dips of AC and pe were compared (Figure 2.2).

43

Figure 2.1. Phenotypic variation between AC and pe. A) Whole plant phenotype of AC and
pe. B & C) Light micrographs of trichomes on leaves of AC and pe. Type VI glandular
trichomes are indicated by arrow heads, scale bar = 2 mm. D) Density of glandular type VI
trichomes on the adaxial leaf surface of AC and pe. Data are presented as the mean of n=8 ± SE
on adaxial side of AC and pe leaves. Asterisks denote significant differences (***, p < 0.001) as
determined by Student’s t tests. E) Comparison of anthocyanin accumulation in the stems of AC
and pe. Stems of the high pigment 1 (hp1) and entirely anthocyaninless (ae) mutants are shown
for comparison. F – I) Autofluorescence of anthocyanins in AC, pe, hp1 and ae visualized by
confocal laser scanning microscopy (Scale bar = 200 μm).

44

Figure 2.2. Comparison of relative monoterpene and sesquiterpene levels in AC and pe
trichomes as determined by leaf dip analysis. Proportions of individual volatile terpenes are
expressed as percentage relative to the abundance in wild type (AC) n=3 ±SE. Asterisks denote
significant differences (*, p< 0.05) as determined by Student’s t tests.

45

Consistent with the reduced abundance of type VI glandular trichomes on the surface of
pe leaves, volatile terpene levels are also significantly reduced to approximately 16 percent of
that observed in wild-type.
Reduced anthocyanin accumulation in pe
To investigate the pale phenotype of pe in more detail, chlorophyll and anthocyanin
levels were determined in fully expanded and meristematic leaves. Chlorophyll levels in AC and
pe are not significantly altered but anthocyanin content is reduced by approximately 85% in pe
(Table 2.1). The anthocyanins represent an endpoint of the phenylpropanoid pathway and are
known to accumulate in epidermal and sub-epidermal cell layers (Martin and Gerats, 1993;
Hichri et al., 2011). The distribution of anthocyanins within the stems of AC and pe was
compared by confocal laser scanning microscopy and stems of the high-pigment 1 and entirely
anthocyaninless (ae) mutants were included as positive and negative controls, respectively
(Figure 2.1E-I). Images reveal two main patterns of anthocyanin distribution in AC, with
accumulation observed in the epidermal and sub-epidermal cells and also surrounding the
vasculature (Figure 2.1F). In contrast anthocyanin accumulation is greatly reduced in stems of pe
(Figure 2.1G).

Table 2.1 Anthocyanin and chlorophyll content of AC and pe leaves
Mature leaves
Anthocyanin content

Meristematic leaves

AC

pe

AC

pe

12.4 ± 2.1***

1.9 ± 0.6

13.1 ± 1.0***

5.4 ± 0.63

54.8 ± 2.0

53.3 ± 3.1

67.3 ± 0.6

66.4 ± 0.8

-1

(AU 535 nm g FW)
Chlorophyll content
-1

(μg ml )
Data represents the mean of n=5 ± SE. Asterisks denote significant differences between genotypes of the same
developmental stage (***, p< 0.001) as determined by Student’s t tests.

46

Lignin content is not altered in pe
Like anthocyanins, lignin also represents an endpoint of the phenylpropanoid pathway
(Vogt, 2010). Reduced lignin synthesis in plants is associated with stunted growth, mediated by
increased salicylic acid levels (Brown et al., 2001; Li et al., 2010; Gallego-Giraldo et al., 2011).
To investigate whether the semi-dwarf phenotype of pe is related to reduced accumulation of
lignin, the total lignin content in leaves and stems of AC and pe was analyzed. The total percent
acetyl bromide soluble lignin (% ABSL) as well as individual lignin monomers of AC and pe
leaves showed no significant differences between the two genotypes (Table 2.2).

-1

Table 2.2. Lignin composition of leaves and stems in AC and pe (μg mg AIR)
Total %ABSL

G

S

H

Leaves AC

2.50 ± 0.16

5.95 ± 0.45

2.20 ± 0.53

0.06 ± 0.01

Leaves pe

2.19 ± 0.23

4.50 ± 0.54

1.75 ± 0.31

0.04 ± 0.01

Stems AC

3.07 ± 0.26

14.57 ± 0.43

2.98 ± 0.24

0.09 ± 0.02

Stems pe

3.06 ± 0.21

14.62 ± 0.45

4.24 ± 0.39

0.09 ± 0.02

AIR, Alcohol Insoluble Residue; ABSL, Acetyl Bromide Soluble Lignin. G, S, and H refer to guaiacyl, syringyl and p
hydroxyphenyl lignin monomers, respectively. Data are shown as means ± SE (n = 5). No significant differences exist
between AC and pe as determined by Student’s t tests.

The pe fruit cuticle has altered physical and chemical properties
Fruits of the pe mutant are highly glossy and are sticky to the touch when compared to
wild type (Figure 2.3A and B) (Butler, 1952). In addition, we observed that ripe pe fruits are
generally crack resistant. The average Young’s elastic modulus (Y) is approximately two-fold
greater than wild type, indicating increased stiffness of the pe fruit peel (Figure 2.3C). Glossiness
and increased stiffness in tomato fruit peels is associated with cutin deficiency (Isaacson et al.,
2009). Scanning electron microscopy (SEM) images of pe fruit indicated reduced cuticle
deposition compared to AC fruit (Figure 2.3D-G).
47

Figure 2.3. Fruit phenotypes of pe. A) Increased glossiness in pe fruits compared to AC. B)
Quantitative analysis of glossiness in ripe fruits of AC and pe. The values of number of pixels
above the saturation threshold represent the mean values (n=5) ±SE. C) Average Young’s
modulus of elasticity (n=5) ±SE. Asterisks denote significant differences (**, p < 0.01; ***, p <
0.001) as determined by Student’s t tests. D – G) Scanning electron micrographs of AC and pe
fruits at the green and ripe stages of development. Note reduced cuticle deposition in the pe
mutant (Scale bar = 20 µm). H & I) Light-micrographs of cuticular lipid distribution in ripe fruits
using Sudan IV staining. Scale bar = 50 µm.
48

Staining of cryosections of the fruit peel of AC and pe with the lipid reactive stain, Sudan IV
confirmed reduced cuticle lipid deposition in the pe mutant (Figure 2.3H-I). The chemical
composition of pe fruit cuticles is significantly altered compared to that of AC fruits. The cutin
monomer load (mass/area) in ripe pe fruits is approximately 2% of that in AC and 6.5% of AC
levels in green fruits (Table 2.3). Furthermore, the large reduction in cutin monomer content is
observed for all hexadecanoic acid-derived cutin monomers, together with cis- and transcoumaric acid, products of the phenylpropanoid pathway. The notable exception to the general
reduction of cutin monomers in pe is the abundance of the hexadecanoic acid itself, which has
slightly increased abundance in mutant fruit, particularly at the green stage of development.
Altered glossiness of plant surfaces is often associated with changes in wax composition (Chen
et al., 2003; Aharoni et al., 2004; Bourdenx et al., 2011). The total wax load is not significantly
different between AC and pe fruits at either the green or ripe stages of development (Table 2.4).
However, the cutin monomer to wax ratio, signifying relative proportion of wax within the cutin
matrix, falls dramatically from approximately 75:1 (1086.5 : 14.42 µg cm-2) in ripe fruits of AC
to 1.6:1 (26.1 : 15.7 µg cm-2) in pe (Table 2.3 and Table 2.4). Also, the composition of
individual wax components is significantly altered. For example, alkanes constitute
approximately 40% of the total wax load in AC fruits but between 55 – 62% in pe fruits due
largely to increases in C31 - C33 alkanes at both the green and ripe stages of fruit development
(Figure 2.3, Tables 2.4 and 2.5).

49

-2

Table 2.3. Cutin monomer composition in fruit cuticles of AC and pe in μg cm (%)
Green fruit

Ripe fruit

Cutin monomer

AC

pe

cis - Coumarate

4.8±1.1(1.6)

ND

6.9±0.8(0.6)

ND

trans - Coumarate

11.3 ± 2.5(3.6)***

0.6±0.1(2.9)

27.6±3.4(2.5)***

0.5±0.1(2.0)

Hexadecanoic acid

0.7±0.1(0.2)

1.82±0.6(9.0)*

1.3±0.2(0.1)

1.4±0.7(5.4)

Hexadecane 1,16 - dioic acid

6.2± 1.0(2.0)

ND

12.7±1.7(1.2)***

0.4±0.1(1.7)

C16:1 Hexadecenoic acid

1.6±0.1(0.5)

ND

9.3±1.3(0.9)***

0.2±0.05(0.8)

16-OH Hexadecanoic acid

4.7±1.1(1.5)***

0.32±0.05(1.6)

22.6±3.9(2.1)***

0.4±0.1(1.6)

9-OH, 16-oxo Hexadecanoic acid

3.9±0.4(1.3)

ND

25.2±5.0(2.3)***

0.4±0.1(1.5)

Octadecane 1,16 - dioic acid

2.1±0.2(0.7)

ND

3.8±0.6(0.4)

ND

8-OH Hexadecane 1,16 - dioic acid

2.5±0.4(0.8)

ND

18.3±4.3(1.7)***

0.6±0.1(2.1)

18-OH Octadecanoic acid

2.3±0.6(0.7)

ND

7.8±1.2(0.7)***

0.3±0.1(1.2)

10, 16-diOH Hexadecanoic acid

176.4±6.0(57.1)***

3.1±1.4(15.5)

827.2±18.5(76.1)***

6.8±0.9(26.0)

9, 18-diOH Octadecanoic acid

2.2±0.6(0.7)

ND

12.6±1.9(1.2)***

0.1±0.05(0.4)

Unidentified

90.0±19.3(29.2)

14.4±1.2(71.0)

111.2±20.5(10.2)

14.9±4.4(57.1)

Total

308.8±20.5***

20.2±1.2

1086.5±22.1***

26.1±5.9

AC

pe

Data represents the mean of n=5 ± SE. ND, not detected. Asterisks denote significant differences between genotypes
of the same developmental stage (*, p < 0.05; ***, p < 0.001) as determined by Student’s t tests.

Table 2.4.Total wax load in fruits of AC and pe at green and ripe stages in µg cm
Total load

Hydrocarbons

Alcohols

Fatty acids

-2

Triterpenoids/
Sterols/
Coumarate

Unidentified

AC green fruit

19.72±1.24

6.86 ± 0.81

1.46 ± 0.06***

0.38 ± 0.05

9.25 ± 0.70***

1.77 ± 0.54

pe green fruit

16.03 ± 2.31

9.81 ± 0.84*

0.53 ± 0.02

0.46 ± 0.07

1.32 ± 0.12

3.91 ± 1.91

AC ripe fruit

14.42 ± 0.91

5.61 ± 0.84

0.96 ± 0.06**

0.74 ± 0.10

4.98 ± 0.21***

2.13 ± 0.25

pe ripe fruit

15.71 ± 1.82

8.86 ± 0.94**

0.64 ± 0.03

1.19 ± 0.13**

2.07 ± 0.17

2.94 ± 0.43

Hydrocarbons in the table include alkanes, iso-alkanes, anteiso alkanes and alkenes. Data are shown as means ± SE
(n = 5). Asterisks denote significant differences between genotypes of the same developmental stage (*, p < 0.05; **,
p < 0.01; ***, p < 0.001) as determined by Student’s t tests.

50

-2

Table 2.5. Fruit wax composition of AC and pe in μg cm x 10

Wax class

C chain
length/compound

Green fruit

Iso alkanes

Anteiso alkanes
Alkenes
Alkanols

Alkenols
Fatty acids

Triterpenoids/
Sterols/Coumarate

C23:0
C25:0
C26:0
C27:0
C28:0
C29:0
C30:0
C31:0
C32:0
C33:0
C29:0
C30:0
C31:0
C32:0
C33:0
C31:0
C32:0
C33:2
C35:2
C20:0
C22:0
C23:0
C24:0
C25:0
C26:0
C27:0
C28:0
C29:0
C30:0
C32:0
C22:2
C24:2
C26:2
C16:0
C18:0
C 20:0
C22:0
C24:0
C25:0
C26:0
trans - Coumarate
psi Taraxasterol
Taraxasterol
delta amyrin
ß amyrin
α amyrin
Cycloartenol
Lupeol
Multiflorenol

Ripe fruit
pe

AC
Alkanes

2

4.6 ± 0.3
1.2 ± 0.3
2.3 ± 0.5
11.4 ± 4.7
3.7 ± 0.5
87.3 ± 4.3***
56.4 ± 4.7
374.6 ± 69.1
45.4 ± 8.7
28.1 ± 6.5
9.1 ± 0.3
0.8 ± 0.3
31.8 ± 2.5
8.8 ± 1.3
13.7 ± 1.2
1.8 ± 0.2
2.8 ± 0.5
2.6 ± 0.8**
ND
10.0 ± 3.1
1.6 ± 0.3
3.3 ± 0.9
1.2 ± 0.3
0.4 ± 0.2
1.8 ± 0.5
3.0 ± 0.3
15.1 ± 1.0
9.4 ± 1.7
27.7 ± 3.0
59.9 ± 3.2
1.3 ± 0.2
11.4 ± 1.7**
0.6 ± 0.2
10.8 ± 0.9
8.0 ± 1.0
12.1 ± 5.5
1.1 ± 0.1
3.2 ± 0.9
0.4 ± 0.2
2.7 ± 0.2
21.3 ± 5.8*
32.9 ± 2.3***
47.9 ± 3.0***
387.6 ± 27.2***
182.7 ± 18.7***
201.3 ± 14.7***
3.6 ± 1.0
8.6 ± 0.9***
40.0 ± 2.7*

3.5 ± 0.4
2.1 ± 1.2
1.1 ± 0.4
34.7 ± 7.2*
3.1 ± 0.8
39.1 ± 5.3
52.5 ± 5.8
508.0 ± 57.4*
101.1 ± 10.7**
118.1 ± 17.1**
8.3 ± 1.1
1.1 ± 0.6
58.4± 4.5***
17.4 ± 1.4**
21.1 ± 1.8**
2.8 ± 1.8
7.3 ± 0.7***
1.1 ± 0.5
ND
5.2 ± 2.5
4.2 ± 1.2
4.8 ± 0.7
1.1 ± 0.3
ND
3.0 ± 0.3
1.5 ± 0.6
12.2 ± 1.8
1.2 ± 0.3
3.4 ± 1.1
11.4 ± 3.4
1.4 ± 0.4
2.2 ± 0.8
1.0 ± 0.4
15.2 ± 2.1
8.4 ± 0.5
4.9 ± 1.1
2.1 ± 1.9
5.9 ± 3.4
5.2 ± 0.4
4.8 ± 2.3
2.7 ± 0.5
5.3 ± 1.7
10.0 ± 1.6
48.1 ± 4.9
27.8 ± 3.7
23.3 ± 2.4
1.4 ± 0.4
1.9 ± 0.3
12.0 ± 8.0

AC
2.1 ± 0.2
1.3 ± 0.6
8.2 ± 1.6
6.2 ± 1.5
3.3 ± 0.5
61.9 ± 3.7
39.8 ± 3.4
276.7 ± 23.4
31.2 ± 4.0
44.1 ± 4.6
9.8 ± 0.5
1.4 ± 0.2
20.2 ± 2.0
5.6 ± 0.6
11.7 ± 1.1
1.0 ± 0.3
1.9 ± 0.4
31.8 ± 8.7**
3.3 ± 0.4
2.7 ± 0.8
1.5 ± 0.3
4.4 ± 0.3
2.9 ± 0.8
4.3 ± 0.8
1.7 ± 0.1
1.7 ± 0.3
15.0 ± 0.9
5.2 ± 1.1**
13.2 ± 0.6***
22.0 ± 3.6***
4.1 ± 1.0
5.3 ± 0.8
12.0 ± 0.5
38.0 ± 1.6
9.6 ± 0.1
7.8 ± 0.4
1.2 ± 0.1
14.8 ± 0.8
1.1 ± 0.2
1.6 ± 0.1
3.0 ± 0.6
15.3 ± 0.7***
25.1 ± 2.3*
211.1 ± 9.5***
105.0 ± 5.3**
112.3 ± 4.3***
1.0 ± 0.2
4.1 ± 0.3
21.0 ± 1.0***

pe
3.2 ± 0.7
1.2 ± 0.2
5.6 ± 0.6
13.0 ± 5.3*
7.5 ± 0.7**
55.1 ± 4.3
58.9 ± 5.6**
415.4 ±42.3**
98.6 ± 6.9***
83.0 ± 13.5*
10.5 ± 1.3
2.4 ± 0.7
75.9 ± 6.7***
23.9 ± 3.0***
21.5 ± 2.6**
1.5 ± 0.3
7.3 ± 0.9***
1.6 ± 0.6
ND
1.1 ± 0.2
3.2 ± 0.3*
7.2 ± 1.2
1.3 ± 0.6
ND
9.6 ± 1.7***
3.5 ± 0.5*
17.1 ± 2.7
1.5 ± 0.2
2.2 ± 0.5
3.1 ± 0.6
1.7 ± 0.3
8.5 ± 1.9
3.6 ± 0.5
76.4 ± 8.0**
15.9 ± 2.0
6.7 ± 1.5
3.7 ± 0.9*
11.2 ± 1.0
2.6 ± 1.8
3.3 ± 0.9
3.4± 0.7
4.3 ± 0.5
11.3 ± 0.6
66.0 ± 4.1
81.4 ± 3.8
31.7 ± 1.8
2.2 ± 0.7
2.6 ± 0.7
4.8 ± 1.2

Data are shown as means ± SE (n = 5). Asterisks denote significant differences between genotypes of the same
developmental stage (*, p < 0.05; **, p < 0.01; ***, p < 0.001) as determined by Student’s t tests.

51

Fatty acids constitute a relatively minor component of the total wax load ranging from 2 5% in both AC and pe and there are no significant differences at the green stage, although ripe
fruits of the mutant have a significantly higher fatty acid content due mainly to a two-fold
increase in hexadecanoic acid. In contrast to the general increase in the levels of long chain
alkanes and fatty acids in the epicuticular wax of pe, the fatty alcohols (alkanols and alkenols)
together with coumarates, sterols and triterpenoids show a general reduction in pe compared to
AC. In particular, the amyrins, which accumulate to similar levels as the alkanes in wild-type
fruit, are reduced by approximately 86% in green fruit of pe (Figure 2.4, Tables 2.4 and 2.5).
Alkanes are well known to constitute much of the epicuticular wax, whereas triterpenoids are
almost exclusively intracuticular waxes (Vogg et al., 2004). Thus the reduction in triterpenoids
but not alkanes with the loss of cutin is consistent with the differential localization of these
waxes.
Altered surface chemistry and increased water conductance in pe leaves
To investigate whether the altered surface chemistry observed in pe is restricted to fruit or
is a more general phenotype of the mutant, the cutin and wax composition of the leaves of AC
and pe were compared. As was observed in fruit, the major monomer identified in tomato leaf
cutin is also 9(10),16-dihydroxyhexadecanoic acid constituting between 20 – 25% of the total
cutin monomer load (Figure 2.5). Additional monomers identified include several fatty acids and
the phenylpropanoids cis- and trans-coumaric acid together with caffeic acid. The overall cutin
monomer load in leaves of pe is 54% of that in AC, with an approximately five-fold reduction in
9(10),16-dihydroxyhexadecanoic acid and significant reductions in additional monomers (Figure
2.5A).

52

Figure 2.4. Relative fruit wax composition in AC and pe. Fruit wax content of green and ripe
stage fruits were analyzed by GC. The wax components were grouped under five classes; i)
Hydrocarbons, consisting of alkanes, branched alkanes and alkenes ii) Alcohols, iii) Fatty acids
iv) Triterpenoids, sterols and coumarates and v) Unidentified compounds. The relative
percentage of each class comprising the total wax load is shown.

53

Tomato leaf cuticular wax components identified in this study comprise n-alkanes, isoalkanes, anteiso alkanes, petacyclic triterpenoids, sterol derivatives and fatty acids. Among these,
alkanes and branched alkanes constituted the major fraction (up to 90%) of all the identified wax
composition in wild-type leaves. The total leaf wax load is reduced by 30% in the mutant. This is
mainly attributed to significant reduction in alkanes. Interestingly all the straight chain alkanes in
the pe mutant were reduced by approximately 50% compared to wild-type, whereas branchedchain alkanes are essentially unaltered (Figure 2.5B). As both wax and cutin loads decrease in
leaf, unlike in fruit (Table 2.3 and Table 2.4), the cutin monomer to wax ratio changes only
fractionally in the pe mutant, from 2.3 (5.85 : 2.53 µg cm-2) to 1.8 (3.20 : 1.80 µg cm-2) (Figure
2.5). This clearly shows a differential regulation of straight chain and branched hydrocarbons in
fruit and leaf cuticular wax, suggesting they may be independently regulated.
Cuticular waxes play a pivotal role in limiting water loss in plants. The levels of alkanes
within cuticular waxes are often positively correlated with increased resistance to cuticular water
loss and an increase in cuticular wax abundance, particularly alkanes can occur during water
stress (Grncarevic and Radler, 1967; Kosma et al., 2009; Bourdenx et al., 2011; Seo et al., 2011).
Furthermore, altered cutin composition may disrupt the intermolecular packing of waxes within
the cutin matrix, although this relationship remains unclear (Pollard et al., 2008; Schreiber, 2010;
Buschhaus and Jetter, 2011). Given the reduction in alkane and cutin monomer content in the
leaves of the pe mutant, the rate of leaf water loss from mutant leaves was determined. The pe
mutant does not have an obviously wilty phenotype however leaf water conductance in intact pe
leaves was approximately 3-fold higher than in AC (Figure 2.6A). Similarly, an increase in the

54

Figure 2.5. Leaf cutin and wax profiles of AC and pe.
55

Figure 2.5 (cont’d)
A) Leaf cutin components were grouped into i) fatty acids and ii) phenylpropanoids. B) Leaf wax
constituents were grouped into four classes i) Alkanes, ii) Branched chain alkanes, iii) Fatty
acids, and iv) Triterpenoids. Inset graphs indicate total cutin and wax loads in AC and pe. Values
represent mean values (n=5) ±SE. Asterisks denote significant differences (*, p < 0.05; **, p <
0.01; ***, p < 0.001) as determined by Student’s t tests.

56

rate of chlorophyll leaching was observed in pe leaves (Figure 2.6B), a phenomena previously
shown to be associated with cuticle permeability (Kosma et al., 2009). Previous research
indicated that defects in cutin biosynthesis can lead to alterations in stomatal structure including
impaired development of cuticular ledges that lie between adjacent guard cells (Li et al., 2007).
Scanning electron micrographs of the leaf surface did not reveal any structural differences in the
stomata of the pe mutant (Figure 2.6C). Furthermore, the average stomatal density on the abaxial
leaf surface in wild type leaves (30 in 0.55 mm2) is higher than in pe leaves (21 in 0.55 mm2)
(Figure 2.6D). Together, these data suggest that enhanced rate of water loss from the leaves of pe
mutant is due to cuticular water loss and not to altered stomatal structure or density.
Root suberin and wax composition are not dramatically altered in pe
The altered cutin and wax composition observed in the leaves and fruits of pe suggested
the possibility of a general perturbation in synthesis of fatty acid derived polymers. This was
investigated by determining the suberin and wax content in roots of pe. The aliphatic
components that account for nearly 90% of the total suberin, together with the aromatic
components are not significantly altered in pe (Figure 2.7A). In addition, the overall root wax
load is largely unaltered in pe although small increases in abundance of β-C22:0 mono-acylglycerol, C29:2 sterol, and C18:0 and C22:0 primary alcohols are observed (Figure 2.7B).

57

Figure 2.6. Increased cuticular permeability in pe. A) Leaf water conductance in AC and pe
(n=5) ±SE. B) Chlorophyll leaching expressed as percent of total leaf chlorophyll extracted after
24 hours (n= 5) ±SE. C) Scanning electron micrographs of representative stomata on the abaxial
leaf surface of AC and pe (Scale bar = 5 µm). D) Stomatal density on abaxial leaf surface area of
0.55mm2 (n= 10 ±SE). Asterisks denote significant differences (*, p < 0.05; ***, p < 0.001) as
determined by Student’s t tests.

58

Figure 2.7. Suberin and root wax composition in AC and pe.
59

Figure 2.7 (cont’d)
A) Suberin monomer content in roots of AC and pe as analyzed by gas chromatography.
Absolute amounts of suberin monomers are shown as mean values (n=5) in ng mg-1 dry weight.
The graph inset indicates the total suberin load. B) Root wax profile in AC and pe. Root waxes
were divided into 6 classes: FFA- free fatty acids; PA- primary alcohols; alkanes; MAG - mono
acyl glycerols; sterols and ferulates (For ferulates, alkyl chain length indicated). Absolute
amounts of individual components are shown as mean values (n=5) in µg mg-1 fresh weight. The
graph inset indicates the total root wax load. Error bars in A & B represent standard error of the
mean. Asterisks denote significant differences (*, p < 0.05; **, p < 0.01) as determined by
Student’s t tests.

60

Mapping of the pe locus and candidate gene identification
The pe locus was provisionally mapped to chromosome 1 of the classical genetic map of
tomato (Mutschler et al., 1987). An F2 mapping population segregating for the pe mutant allele
was generated through crosses between S. lycopersicum (pe/pe) (LA2467) and S.
pimpinellifolium (PE/PE) (LA1589). The mutant plants were pale green in color and this
phenotype co-segregated with the cutin deficient, sticky and glossy fruit phenotype in the F2
population. The F2 population was genotyped with chromosome 1 molecular markers, revealing
that the pe locus is located within a 424 kb interval between C2_At4g00090 and cTOA-13-J3
(Figure 2.8A). This mapping interval contains the CUTIN DEFICIENT 2 (CD2) gene, which
encodes a HD-ZIP IV protein, with homology to ANL2 of Arabidopsis (Kubo et al., 1999;
Isaacson et al., 2009). A single G > A substitution causes the conversion of a conserved glycine
to arginine at position 736 of the protein at the cd2 locus which results in a glossy and cutin
deficient fruit phenotype with a total cutin load of approximately 10% of wild type fruits
(Isaacson et al., 2009). The similarity in phenotype and map position of pe and cd2 suggested
that pe may be allelic to cd2. To investigate this hypothesis, the CD2 gene was cloned and
sequenced from the pe mutant revealing the insertion of a G nucleotide following nucleotide
2045. This insertion causes a frame shift mutation resulting in the incorporation of four spurious
amino acids followed by a premature stop codon that truncates the predicted protein by 160
amino acids in pe (Figure 2.8A). To confirm that a mutation in the CD2 gene is responsible for
the pe mutant phenotype, VIGS experiments were performed using two constructs targeting
separate regions of the CD2 gene. CD2 silenced lines showed a 20-30% reduction in height
compared with TRV2-empty vector control lines and leaves showed sectors of pale coloration
characteristic of the pe mutant (Figure 2.8B-D).

61

Figure 2.8. Characterization of the pe locus and silencing of CD2.
62

Figure 2.8 (cont’d)
A) Genetic map of the pe locus. Genetic markers and the number of recombinant individuals
between adjacent markers from a total of 114 F2 plants are shown. The approximate physical
distance between the flanking markers and the pe locus is indicated. Sequence analysis of the
CD2 gene from AC and pe revealed a single G nucleotide insertion following nucleotide 2045
leading to a frame shift mutation. B – G) Silencing of CD2 recreates the pe mutant phenotype. B)
Pale leaf phenotype in a CD2-2 VIGS line. C) Short stature of a CD2-2 VIGS line D) Plant
height of CD2 VIGS lines. E) Anthocyanin content of CD2 VIGS lines. F) The major leaf cutin
monomer 9(10), 16-dihydroxyhexadecanoic acid is reduced in CD2 silenced lines. G) CD2
silencing impairs leaf cuticle barrier properties and increases water conductivity. All values D –
G represent mean values (n=5) ±SE. TRV2 empty vector control lines were used for all
comparisons. Asterisks denote significant differences (*, p < 0.05; ***, p < 0.001) as determined
by Student’s t tests.

63

Anthocyanin levels were reduced in the CD2 silenced lines by approximately 30-35%
(Figure 2.8E). The major leaf cutin monomer 9(10), 16-dihydroxyhexadecanoic acid was
reduced by 50% in the silenced lines and there was a concomitant increase in the leaf water
conductance (Figure 2.8F and G). Together, these data indicate that the pe mutant phenotype is
caused by a mutation in CD2.
Mutation at the anthocyaninless 2 locus leads to cutin deficiency
Phylogenetic analysis indicated that CD2 is closely related to a number of HD-ZIP IV
proteins including several that impact epidermal cell development and cuticle biosynthesis
(Nakamura et al., 2006; Javelle et al., 2010; Javelle et al., 2011) (Figure 2.9A). HD-ZIP IV genes
are typically expressed in epidermal cells and expression analysis indicated that CD2 transcripts
are enriched in stem peels, which are comprised of a mixture of epidermal and sub-epidermal
cells, compared to the levels observed in whole stem and stem core with the peel removed
(Figure 2.9B).
ANL2 is the closest Arabidopsis homolog to CD2 (Figure 2.9A) and is also preferentially
expressed in epidermal cells (http://efp.ucr.edu/cgi-bin/absolute.cgi). Furthermore, both pe and
anl2 mutants have reduced anthocyanin accumulation (Kubo et al., 1999) (Figure 2.1). These
similarities led us to hypothesize that anl2 mutants would also be cutin deficient. This was
confirmed in rosette leaves of anl2, which exhibit approximately a 40% reduction in cutin
monomer load together with a 25% reduction in the alkane load of the cuticular waxes compared
to Col-0 (Figure 2.10).

64

Figure 2.9. Phylogenetic and expression analysis of CD2.
65

Figure 2.9 (cont’d)
A) A neighbor-joining phylogenetic tree derived from a multiple sequence alignment of the
deduced full-length amino acid sequences of CD2 homologs constructed using MEGA V5.0
software (Tamura et al., 2011). HDG7 is included as an out-group. Bootstrap values are from
2,000 replicates, indicated above the nodes. Proteins highlighted in bold have defined functions
in cuticle biosynthesis. B) qRT-PCR analysis of CD2 expression in stem peel of AC relative to
stem core and whole stem. Three biological and three technical replications for each sample were
analyzed. Data are presented as the mean expression values ±SE relative to that observed in
whole stem. Values labeled with different letter are significantly different (LS Means, p < 0.05).

66

Figure 2.10. Cutin monomer and wax profile of Arabidopsis Col-0 and anl2 leaves. A) Leaf
cutin monomer composition (DCA, α, ω dicarboxylic acid). B) Alkane constituents of leaf wax.
Inset graphs indicate total cutin and alkane load. Data represents mean values (n=5) ±SE.
Asterisks denote significant differences (*, p < 0.05; **, p < 0.01; ***, p < 0.001) as determined
by Student’s t tests.
67

Discussion
Characterization of pe identifies additional roles for CD2
Phenotypic characterization indicates that the pe mutant has altered fruit surface
chemistry leading to reduced cutin levels and a modified surface wax composition (Figure 2.3,
Table 2.3 and Table 2.4). The altered chemical profile leads to a change in the physical
properties of the fruit peel including increased glossiness and a higher Young’s elastic modulus
(Figure 2.3A-C). These phenotypes are identical to those previously described in cutin deficient
mutants of tomato where cutin deficiency was correlated with altered biomechanical and
structural properties together with enhanced susceptibility to microbial infection (Isaacson et al.,
2009). A combination of genetic mapping and gene cloning revealed that pe represents a new
mutant allele of CD2 that truncates the predicted protein by 160 amino acids and therefore likely
represents a null allele (Figure 2.3A).
Previously, the cd2 phenotype was associated with fruit-specific alteration in cuticle
biosynthesis (Isaacson et al., 2009). However, as shown in this study, many additional
phenotypes are evident in pe including a semi-dwarf stature, reduced anthocyanin accumulation,
altered cutin and wax deposition in leaves, increased permeability of the leaf cuticle, and reduced
trichome density together with lower volatile terpene production (Figure 2.1, Figure 2.2, Figure
2.5, Figure 2.6 and Table 2.1). These phenotypes were confirmed in the cd2 allele and VIGS
lines targeting CD2 (Figure 2.8 and Figure 2.11 and Figure 2.12). Together, these data reveal a
broad role for CD2 in plant development, and particularly in relation to metabolism within
epidermal and sub-epidermal cells, which is supported by the epidermal enriched expression
pattern of CD2 in stems (Figure 2.9B) and in tomato fruit (Matas et al., 2011).

68

Figure 2.11. Comparison of leaf cutin and wax composition in M82 and cd2.

69

Figure 2.11 (cont’d)
A) Leaf cutin monomers that are identified and quantified. B) Leaf wax constituents were
grouped into four classes i) Alkanes ii) Branched chain alkanes iii) Fatty acids iv) Triterpenoids.
Inset graphs indicate total cutin and wax loads in M82 and cd2. Values represent mean values
(n=5) ±SE. Asterisks denote significant differences (*, p < 0.05; **, p < 0.01; ***, p < 0.001) as
determined by Student’s t tests.

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Figure 2.12. Trichome density on leaves and anthocyanin accumulation in leaves and stem
peels of A) M82 and B) cd2. Scale bars represent 2 mm. C, Density of type VI glandular
trichomes on the adaxial leaf surface of M82 and cd2. D, Anthocyanin contents (AU 535 nm) of
M82 and cd2 leaves and stem peels. The data shows the mean trichome number (n=5 ±SE).
Asterisks denote significant difference ( **, p<0.01) as determined by student’s t test.

71

HD-ZIP IV proteins and cuticle biosynthesis
CD2 belongs to a subgroup of HD-ZIP IV proteins, several of which have defined roles
in epidermal cell development and are preferentially expressed within the epidermis (Figure
2.9B) (Javelle et al., 2011). Proteins of this family are characterized by HD-ZIP domain followed
by a steroidogenic acute regulatory lipid transfer (START) domain. The fact that HD-ZIP IV
proteins are widely distributed in lower plants, gymnosperms and angiosperms but are absent in
algae is consistent with a role in the evolution of land plants (Ponting and Aravind, 1999;
Mukherjee et al., 2009). Several closely related CD2 homologs influence cuticle composition
and/or regulate genes involved in cuticle biosynthesis. For example, OUTER CELL LAYER 1
(OCL1) of maize is expressed early in embryo development, prior to protoderm formation and is
subsequently expressed in the L1 cell layer in the shoot apical meristem (Ingram et al., 1999).
Over-expression of OCL1 in maize altered the wax composition in leaf blade as well as sheath
and increased the expression of several genes involved in wax biosynthesis and transport (Javelle
et al., 2010). Furthermore, transient assays suggested that activation of the expression of a wax
transporter and a non-specific lipid binding protein likely occurs through direct binding of OCL1
to regulatory regions within these genes (Javelle et al., 2010). Similarly, Arabidopsis HDG1
directly regulates the expression of BODYGUARD (BDG) and FIDDLEHEAD, two genes
involved in cuticle biosynthesis and disruption of HDG1 function leads to cuticles with increased
permeability (Wu et al., 2011). Furthermore, Arabidopsis ANL2 and HDG1 are preferentially
expressed in epidermal or sub-epidermal cells (Suh et al., 2005; Kubo et al., 2008)
(http://efp.ucr.edu/cgi-bin/absolute.cgi) and our analysis indicates that anl2 has a cutin deficient
phenotype (Figure 2.10).

72

Overall, the characterized members of the CD2 subclade all possess a common role in the
regulation of cuticle biosynthesis, even though the composition of cutin can vary between
species and tissues. For example, while tomato cutin is predominantly comprised of
hexadecanoic acid-derived monomers, Arabidopsis leaves have a C18:2 dicarboxylic acid-rich
cutin (Table 2.3 and Figures. 2.5 and 2.10). These cutins require different sets of biosynthetic
genes (Li et al., 2007; Pollard et al., 2008; Li-Beisson et al., 2009) suggesting that while the
regulation of cuticle biosynthesis may be generally conserved, these regulatory proteins may
target distinct sets of biosynthetic genes in diverse species or tissues contributing to their
different cuticle composition. Furthermore, given that several members of the CD2 subclade
influence cuticle biosynthesis (Figure 2.9A), it is possible that the phylogenetically closelyrelated and as yet uncharacterized genes, may also have a similar role.
A role for CD2 in reducing leaf water loss
The cuticle regulates non-stomatal water loss and the amount and composition of
cuticular wax is associated with cuticle permeability to water. For example, mutants or
transgenic lines with reduced levels of very long chain alkanes often display elevated rates of
water loss (Vogg et al., 2004; Leide et al., 2007; Qin et al., 2011). Furthermore, enhancing the
levels of these wax components can restrict water loss leading to improved drought tolerance
(Bourdenx et al., 2011; Seo et al., 2011). Similarly, drought-stressed plants can increase cuticular
wax biosynthesis as a defense mechanism to reduce further water loss (Kosma et al., 2009).
The wax composition of leaves and fruits vary in pe when compared to AC. For example,
hydrocarbons are elevated in pe fruits whereas in leaves the straight-chain alkanes are
significantly reduced compared to wild-type (Figure 2.5B and Table 2.5). The same trend for
cutin and wax is observed in the fruit (Isaacson et al., 2009) and leaves (Figure 2.11) of cd2
73

when compared to the M82 parental line. The basis for the differential CD2-mediated control of
wax accumulation in leaves and fruits is currently unclear but may occur through altered gene
regulation between these tissues or possibly altered fluxes of precursors in mutant tissues. For
example, a proportion of the hexadecanoic acid precursor, typically utilized in cutin biosynthesis
may be channeled toward alkane biosynthesis when cuticle biosynthesis is limited in pe and cd2
fruits. However, this is unlikely to be a general phenomenon as an increase in alkane
accumulation is not observed in cd1 and cd3 fruits (Isaacson et al., 2009).
Our data suggest a correlation between cuticular wax composition in pe and leaf cuticle
permeability. For example, leaf water conductance and chlorophyll leaching are increased in pe
leaves compared to AC (Figure 2.6). A similar trend was observed in CD2 silenced lines (Figure
2.8G). In contrast, fruits of the pe and cd2 mutants, which have higher levels of long chain
alkanes, do not exhibit enhanced rates of post-harvest water loss (Table 2.5) (Isaacson et al.,
2009). Although disruption of individual cutin or wax components can alter permeability, the
physical consequences of these changes on the structure of the cutin matrix, is not well
understood. Alkanes are proposed to influence cuticle permeability by forming waterimpermeable crystalline regions within the cuticle and the packaging of the wax crystals is likely
dependent upon the cutin structure (Pollard et al., 2008; Buschhaus and Jetter, 2011). Therefore,
while it is not possible to more exactly define the underlying reason for the increased
permeability of pe leaves, it could be due to one, or a combination of the following factors,
including decreased alkane levels, a decreased cutin load, an alteration in the cutin to wax ratio,
or an as yet undefined biochemical change. A comparison of wax biosynthesis between wildtype and pe leaves and fruit tissues, together with a more in depth understanding of wax

74

deposition in these genotypes and tissues, may provide insight into how CD2 influences cuticular
permeability.
Leaf water loss is also intricately linked to stomatal structure and physiology and several
lines of research have highlighted possible links between cuticle composition and stomatal
density and structure. Cutin deficiency in Arabidopsis leads to altered stomatal morphology
including a reduction in the cuticular ledges (Li et al., 2007). No structural alterations were
observed in stomata of the pe mutant suggesting that CD2 may not influence cuticle biosynthesis
in leaf guard cells (Figure 2.6C). Similarly, previous reports have shown that altered cuticular
wax composition can positively or negatively influence stomatal density in Arabidopsis (Gray et
al., 2000; Chen et al., 2003; Yang et al., 2011). The exact mechanisms through which this
process occurs are unknown although it is possible that the altered composition of leaf cuticular
waxes in pe may exert a compensatory influence leading to the observed reduction in stomatal
density (Figure 2.6D).
CD2 and phenylpropanoid biosynthesis
Flux into the phenylpropanoid pathway through the enzyme phenylalanine ammonia
lyase represents a branch point between primary and specialized metabolism in plants. The
phenylpropanoid pathway synthesizes a wide range of compounds including flavonols,
anthocyanins, lignins, suberin and cutin aromatics, and tannins, which have diverse roles in
plants as structural polymers, pigments, UV protectants and signaling molecules (Hahlbrock and
Scheel, 1989; Vogt, 2010). In agreement with the epidermal enriched expression pattern of CD2
(Figure 2.9B), the pe mutant exhibits reduced accumulation of a subset of phenylpropanoids that
typically are synthesized and accumulate in the epidermal and sub-epidermal cells. In addition to
the anthocyanins, which are severely reduced in pe tissues, coumarates, which comprise between
75

3 and 5% of the total cutin monomer load in wild type tomato fruits, are also reduced in pe
(Figure 2.1, Table 2.1 and Table 2.3). The reduction in coumarate is readily ascribed to the
reduction of the cutin aliphatic hydroxy fatty acid monomers, to which coumarate is likely
esterified. In contrast, the abundance of lignin and suberin ferulates, two biopolymers derived
from phenylpropanoid precursors that are not typically associated with epidermal cells, are
unaltered in pe (Figure 2.7 and Table 2.2). It will be of interest to determine if additional
phenylpropanoid-derived metabolites are altered in the epidermal cells of pe and to understand
the flux through the pathway in these cells in the pe mutant. In addition, the role of CD2/ANL2 in
directly regulating anthocyanin biosynthesis, as well as understanding their relationship to
known transcriptional regulators of anthocyanin biosynthesis warrants additional investigation.
CD2 as a regulator of epidermal cell function
HD-ZIP IV proteins are master regulators of epidermal cell fate and function in higher
plants and have roles in cuticle biosynthesis together with patterning of trichome and stomata
(Javelle et al., 2011). Our biochemical characterization of the pe mutant of tomato has revealed
alterations in multiple pathways associated with epidermal cell function that have evolved to
facilitate plant adaptation to stress, including altered cuticle biosynthesis in leaves and fruits,
reduced anthocyanin accumulation, and lower trichome and stomatal densities. The subsequent
identification of pe as an allele of CD2 extends current knowledge of the role of CD2 beyond
cuticle biosynthesis in tomato fruit to include a spectrum of altered phenotypes associated with
epidermal cell function. While mutant alleles of CD2 influence a broad range of epidermal cell
phenotypes, it is difficult to determine whether these alterations constitute primary or secondary
effects. For example, altered stomatal and trichome densities can result from perturbation of
cuticular wax biosynthesis, including mutations that affect catalytic enzymes, suggesting that
76

disruption of cuticle structure and / or physiology induces secondary phenotypes (Gray et al.,
2000; Chen et al., 2003; Kurata et al., 2003; Aharoni et al., 2004). Establishing cause and effect
relationships with respect to the altered stomatal and trichome density phenotypes observed in pe
will be complex.
Our data suggests that CD2 is a master regulator of epidermal cell function in tomato.
Although the exact role of CD2 in specifying epidermis-related phenotypes remains to be
defined, it is possible that CD2 directly regulates the expression of the biosynthetic genes
involved in synthesis of diverse epidermal cell associated molecules. Alternatively, it is equally
probable that CD2 acts upstream of these pathways, within a transcriptional network that
maintains epidermal cell identity and function. Defining the immediate targets of CD2 will
provide insight into these questions and will help identify the regulatory networks that control
specialized metabolism within plant epidermal cells.

77

Materials and Methods
Plant material and growth conditions
Seeds of the sticky peel (pe/pe) (LA2467), high pigment-1 (hp-1/hp-1) (LA3538), and
entirely anthocyaninless (ae/ae) (LA3612) mutants, together with S. pimpinellifolium (LA1589)
were obtained from the Tomato Genetics Resource Center, UC Davis. The cultivar AC was
originally obtained from the Glasshouse Crops Research Institute (Littlehampton, Sussex, UK).
As the genetic background of LA2467 was not known, a segregating BC 2F2 population was
generated from an initial cross of LA2467 (pe/pe) with AC (PE/PE) as the recurrent parent. The
mutant plants (pe/pe) were selected based on phenotype; short stature, pale green color of the
stem and leaves and stickiness of the fruit surface, all of which co-segregated. This population
was used for all experiments. Unless otherwise stated, plants were grown in peat-based compost
supplemented with fertilizer in greenhouses at Michigan State University under 16 h day (25°C)
and 8 h night (20°C). Arabidopsis seeds of anl2 (SALK_000196) were described previously
(Nakamura et al., 2006)

and sown in soil (1:1:1 Sure mix: Medium vermiculite: Perlite,

Michigan Grower Products Inc., www.suremix.com) and after 3d at 4°C, maintained in a growth
chamber under a 16 h photoperiod (145 µmol m-2 s-1) at 22°C at a relative humidity of 65%.
Tomato fruits were harvested when they reached 35 to 40 mm diameter for green stage or at
seven days after the onset of ripening (breaker +7 days) for ripe stage of development. All
experiments involving leaves, stems and roots were performed on five-week-old plants unless
otherwise stated. Rosette leaves from seven-week-old Arabidopsis plants were used for cutin
analysis.

78

Extraction and analysis of cuticular lipids
Extraction of surface waxes from leaf, root and fruit tissues of tomato or from leaves of
Arabidopsis was accomplished through dipping tissue samples in chloroform containing 1%
methanol for 2 minutes in the case of fruits and 1 min for other tissues. The surface areas of
leaves and fruits were determined prior to wax extraction using a LI-3100C Area Meter (Li-Cor,
http://www.licor.com) and Tomato Analyzer version 2.1.0.0., respectively (Brewer et al., 2006).
Internal standards of tetracosane, hexadecanol and heptadecanoic acid (5μg/ml each) and ωpentadecalactone,

methyl

heptadecanoate

(10

μg/ml

each)

(Sigma-Aldrich,

http://www.sigmaaldrich.com) were added to each wax and cutin extract, respectively.
Preparation of tri-methyl-silyl derivatives of wax components, followed by delipidation, basecatalyzed trans-methylation and acetyl derivatization of cutin monomers from all the tissues was
performed as previously described (Molina et al., 2006). GC (FID) (6890N, Agilent
technologies) analysis of the derivatized wax or fatty acid methyl ester products used a DB-5
capillary column. GC temperature was programmed from 140°C to 310°C for cutin and to 330°C
for wax, at 5°C min-1. Samples were injected in split mode (330°C injector temperature).
Quantification was based on FID ion current using peak areas relative to internal standard peak
areas. When necessary, compound identification was performed by GC-MS (EI mode) using an
Agilent Technologies 6850 GC, equipped with an HP-5 MS column connected to an Agilent
5975 mass spectrometer. Helium was used as carrier gas at 2 ml min-1 and oven temperature was
programmed from 120°C to 340°C at 10°C min-1. Split-less injection was used and ions were
collected in scan mode (40-800 atomic mass units) with peaks quantified on the basis of their
total ion current.

79

Imaging of cuticular lipids and scanning electron microscopy
Cryosections of fruit pericarp, 8µm thick, were generated using a Microm HM550
cryostat (Thermo scientific, http://www.thermoscientific.com) and stored at -20°C. Sections
were stained using Sudan IV, 0.1% w/v in isopropyl alcohol. Bright field images were collected
using an Olympus IX81 inverted microscope (Olympus, www.olympus.com) configured with
UPlan SApo 20X objective, NA0.75. The images were captured using an Olympus DP72 color
camera and analyzed with DP2-BSW software. For imaging of fruit cuticles by scanning electron
microscopy, fruits were hand-sectioned and fixed for 30 min in 4% glutaraldehyde followed by
30 min incubation in 0.1 M phosphate buffer (pH 7.4). Further processing of samples and
imaging are according to (Li et al., 2007). The same procedures were followed for collecting leaf
stomatal images.
Imaging anthocyanin distribution
Images of hand-cut cross sections of stem tissue were collected using an Olympus
FluoView FV1000 Confocal Laser Scanning Microscope (Olympus) configured on IX81
inverted microscope with a 10X UPLSAPO objective, NA0.4. An excitation wavelength of 559
nm was generated using solid state laser, while the fluorescence emission was collected using a
570 to 640 nm band pass filter, to eliminate the detection of chlorophyll fluorescence. The DIC
transmitted laser light image was collected using the 559 nm laser line and the images presented
as overlays of confocal and DIC images.
Determination of stomatal and trichome densities
For stomatal density measurements, leaf surface imprints on abaxial sides of the leaf
surface were taken using transparent nail polish and placed onto microscope slides. Imaging was

80

performed using a Zeiss Axiophot microscope (Carl Zeiss, Inc., http://www.zeiss.com) at 20X,
image processing was accomplished using Image-pro plus image analysis software (Media
Cybernetics, http://www.mediacy.com). Stomatal density was expressed as number of stomata in
five different fields of 0.55 mm2 per leaflet of five individual leaflets. Trichome density was
analyzed on three week-old plants of AC, M82, pe and cd2 using plant growth conditions and
methods previously described (Kang et al., 2010).
Analysis of volatile terpenes from tomato trichomes
Volatile terpene levels were determined in three-week old plants. Briefly, a leaflet from
the second newly emerging leaf was dipped in 1 ml of methyl tert-butyl ether containing 5 ng/ l
of tetradecane as an internal standard, and allowed to rock for 1 min. GC-MS analysis was
performed as previously described (Schilmiller et al., 2009). Terpene identification was based on
comparison of mass spectra and retention times with those available in an essential oil library
(Adams, 2009).
Determination of fruit glossiness
The glossiness of red-ripe fruits was measured using a custom made gonioreflectometer
consisting of a Fire-iTM digital camera (http://www.unibrain.com) and two 60 W incandescent
light bulbs located in a light impervious container. Thirty images were taken for each fruit, as it
rotated 360°. The average number of saturated blue pixels for all the images was recorded as an
index of fruit gloss measured in gloss units (A gloss unit is defined as number of pixels above the
saturation threshold which is fixed based on specular reflectance of the sample image)
programmed using MATLAB (MathWorks , www.mathworks.com). The instrument was
calibrated using standard glossy and non-glossy green and red spheres prior to data collection.

81

Mechanical analysis of the fruit cuticle
Young’s elastic modulus (Y) of isolated tomato cuticles of 3 cm X 0.5 cm dimension was
measured

using

a

dynamic

mechanical

analyzer

(DMA)

Q800

(TA

Instruments

http://www.tainstruments.com) in constant stress ramp mode at room temperature until breakage
was observed. Y values were calculated using Universal Analysis 2000 software (TA
Instruments).
Determination of anthocyanin and chlorophyll content
Anthocyanins were extracted from leaves according to (Peters et al., 1998). The
absorbance (A535) of the aqueous phase was determined spectrophotometrically using a Hitachi
U3000 spectrophotometer (Hitachi High Technologies America, Inc, http://www.hitachihta.com). Chlorophyll content was sampled from 3 leaf discs. Discs were incubated in darkness
for 48 h at 4°C in 100% DMSO. A647 and A664 were determined spectrophotometrically as
described above. Chlorophyll a and b were calculated according to (Moran, 1982).
Determination of leaf epidermal permeability
Leaf water conductance was measured using Li-Cor 6400 (Li-Cor). Plants were dark
acclimated for 3 h prior to measurements being made to eliminate the effect of stomatal
conductance. Chlorophyll leaching assays were performed on fully expanded leaves. The amount
of chlorophyll extracted into the solution was quantified every 30 minutes as described above
and data was expressed as a percentage of the total chlorophyll extracted after 24 h according to
(Kosma et al., 2009).

82

Lignin analysis
Lignin was extracted as previously described (Foster et al., 2010). Results are reported as
% acetyl bromide soluble lignin (ABSL) based on the dry weight of the alcohol insoluble residue
according to (York et al., 1986; Fukushima et al., 1991; Robinson and Mansfield, 2009). For the
calculation of % ABSL content of tomato, the molar extinction coefficient for Arabidopsis
(23.35 g-1l cm-1 ) was used as an estimate given that no tomato-specific coefficients were
available at the time of analysis (Chang et al., 2008).
Genetic mapping
An inter-specific F2 population of 114 individuals derived from a cross between S.
lycopersicum (pe/pe) (LA2467) and S. pimpinellifolium (PE/PE) (LA1589), was phenotyped for
paleness of the vegetative tissues and stickiness of the fruit peel and simultaneously genotyped
with chromosome 1 molecular markers, previously selected to be polymorphic between LA2467
and LA1589 (Table 2.6). Genomic DNA was extracted from expanding leaves as previously
described (Barry et al., 2005). Details of genetic maps and molecular markers can be accessed
through the Sol Genomics Network (http://solgenomics.net).
Cloning of CD2 and phylogenetic analysis
Total RNA was extracted from expanding leaves of AC and pe using the RNeasy Mini
Kit (Qiagen, http://www.qiagen.com). 1μg of RNA was used as a template for oligo dT primed
first strand cDNA synthesis using the Transcriptor First Strand cDNA synthesis kit (Roche
Applied Science, https://www.roche-applied-science.com). The coding region of CD2 was
amplified

from

cDNA

using

Pfu

Ultra

DNA

polymerase

(Agilent

technologies,

http://www.home.agilent.com) using the primers CD2-F and CD2-R (Table 2.7). PCR fragments

83

were purified using the PureLink PCR purification kit (Invitrogen, http://www.invitrogen.com)
and cloned into the pCR™4Blunt-TOPO

vector (Invitrogen). Sequencing of inserts was

accomplished using vector primers and the gene-specific primers CD1 through CD6 (Table 2.7).
Sequences were assembled using Sequencher 4.7 software (Gene Codes Corporation,
http://genecodes.com). A neighbor-joining phylogenetic tree was constructed from a multiple
sequence alignment of the deduced full-length amino acid sequences of selected CD2 homologs
using MEGA V5.0 software (Table 2.8) (Tamura et al., 2011).
Construct assembly and silencing of CD2
Two constructs, CD2V1 and CD2V2, were assembled in the TRV2-LIC vector as
previously described (Dong et al., 2007) using the primers CD2V1-F, CD2V1-R, CD2V2-F and
CD2V2-R (Table 2.7). VIGS experiments were performed as described by (Velasquez et al.,
2009). Tomato cotyledons were infiltrated with Agrobacterium tumefaciens strain GV3101
cultures before the appearance of the first true leaves and tissues were harvested four weeks after
infiltration. TRV2-LIC empty vector plants were used as controls and TRV2::PHYTOENE
DESATURASE plants were used to monitor the progression of gene silencing. The VIGS plants
were maintained in a growth chamber under a 16 h photoperiod (145 µmol m-2 s-1) at a constant
temperature of 27°C and a relative humidity of 60%.
qRT-PCR analysis
For CD2 expression analysis stems were divided into stem peel, stem core (stem devoid
of peel) and whole stem. Total RNA was isolated as described above. RNA was treated on
column with DNase (Qiagen) and 1µg of RNA was used for reverse transcription using
SuperScriptTM III First-Strand Synthesis System (Invitrogen). Gene-specific primers for the

84

analyzed genes were designed by Primer Express 3.0 (ABI); (listed in Table 2.7). The PCR
reactions

were

performed

with

FAST

SYBR

Master

Mix,

2X

(ABI,

http://www.appliedbiosystems.com) in a 25 µL volume using an Applied Biosystems
StepOnePlusTM Real-Time PCR System (ABI) with the following cycling program: 10 min at
95°C, 15 s at 95°C followed by 40 cycles of 1 min at 60°C, 15 s at 95°C, 1 min at 60°C. The
comparative ΔΔCT method was performed according to (Balaji et al., 2008).
Statistical analyses
Statistical analyses were performed using SAS (SAS Institute, www.sas.com). The
genotypic constituents were evaluated by Student’s t-test and LS Means.
Acknowledgements
We thank Carol Flegler, and Drs. Melinda Frame and Alicia Pastor (Centre for Advanced
Microscopy, Michigan State University) for assistance with microscopy. We are grateful to Dr.
Renfu Lu (Department of Biosystems and Agricultural Engineering, Michigan State University)
for help with glossiness measurements; Clifton Foster and the Analytical Cell Wall Facility
(Great Lakes Bio-Energy Research Center, Michigan State University) for lignin analysis; Dr.
Michael Rich (Composite Materials and Structure Center, College of Engineering, Michigan
State University) for guidance with dynamic mechanical analyzer experiments; and Dr. Eliana
Gonzales-Vigil (Department of Horticulture, Michigan State University) for help with analysis of
trichome chemistry. We also thank Dr. Jocelyn Rose (Cornell University) and Dr. Taku
Takahashi (Okayama University) for providing seeds of the cd2 allele and anl2 mutant,
respectively.

85

Table 2.6. PCR based genetic markers flanking the pe locus
Marker
C2_At5g13700

C2_At4g00090

cTOA-13-J3

U151020

C2_At2g38730

C2_At1g56345

C2_At5g49880

Primers
For

5'-TCATTCTTCATGATTTTGAAATGGC- 3'

Rev

5'- AGTCGACTGTCCATAATTTTACCCTC-3'

For

5'- AGATATTGGCCACCACTCATGGTTC-3'

Rev

5'- AGGCGACCATGCCATGTCCG-3'

For

5'- AGTGATGTCGAGGGTCCTCTGAAAAC -3'

Rev

5'- GTACAAATAGGCCTATCAACAACA -3'

For

5'- GTAATGAAGAATCTGGCTCTGCAA -3'

Rev

5'-GTCCAATTTAATTTCACTCGATGA -3'

For

5'- AGCGGACCAAACACTAATGGATG-3'

Rev

5'- AGCCACATTCTCAATCTTCCTGAC-3'

For

5'- TCGGTGGCAACCACAGATTACCCAG-3'

Rev

5'- ATCGAGTCTATTAGCTAAATGAAGCTC-3'

For

5'-TCGGCTCCGTTGCCGGAATC-3'

Rev

Restriction enzyme
HhaI

RsaI

RsaI

DdeI

AluI

MspI

RsaI

5'- ACTGCTTAAGGCATCAAAAAACTC-3'

Restriction enzyme used to resolve polymorphisms between S. lycopersicum LA2467 (pe/pe) and S. pimpinellifolium
LA1589 (PE/PE).

86

Table 2.7. Oligonucleotide primers used in the study
Gene
CD2
#GQ222185

Primer

Use

CD2-F
CD2-R

5’-GATACAGTGAATCTGTCAAATATAC-3
5’-TTAGCTTTCGCATTGAAGTGCAGCT-3

C&S

CD2-SEQ1

5’-GATCGAGTTTGTGCACTTGCTGGC -3

C&S

CD2-SEQ2

5’-ATCAAGTGGTATGGGTGGAACCA-3

C&S

CD2-SEQ3

5’-TGGTGCACCAGAAGTTTCACGGA-3

C&S

CD2-SEQ4

5’- GCCAGTGTGAGTGTCTTGCAATTC-3

C&S

CD2-SEQ5

5’-TGTTGTTGGTACATTACTCATACC-3

C&S

CD2-SEQ6

5’-GCAAATCCAAGAACTTGAATCTC-3

C&S

CD2-SEQ7

5’-GTTGAAGCACAAACACCAGCACA-3

C&S

C&S

CD2V1-F

5’-CGACGACAAGACCCTGCAAGAGAATGATAAGCTTCGTGC-3

VIGS

CD2V1-R

5’-GAGGAGAAGAGCCCTGCAGCCAAAGCAAGTTCAAGATAC-3

VIGS

CD2V2-F

5’-CGACGACAAGACCCTGTCATCCACCGTGTCTGCCAGGGA -3

VIGS

CD2V2-R

5’-GAGGAGAAGAGCCCTGGAATATCAACTGGCGCGTATACA-3

CD2Q-F
GAPDH
#U97257

Sequence

VIGS

5’-TGGGTGGAACCAGAAATGGT-3

qRT-PCR

CD2Q-R

5’-TGGTGAAAGCACTTGGAGTTCA-3

qRT-PCR

GAPDHQ-F

5’-ATGCTCCCATGTTTGTTGTGGGTG-3

qRT-PCR

GAPDHQ-R

5’-TTAGCCAAAGGTGCAAGGCAGTTC-3

qRT-PCR

(Balaji et al.,
2008)

Cloning & Sequencing – C&S; VIGS constructs – VIGS
Balaji, V., Mayrose, M., Sherf, O., Jacob-Hirsch, J., Eichenlaub, R., Iraki, N., Manulis-Sasson, S., Rechavi, G.,
Barash, I. and Sessa, G. (2008) Tomato transcriptional changes in response to Clavibacter michiganensis subsp.
michiganensis reveal a role for ethylene in disease development. Plant Physiol., 146, 1797-1809.

87

Table 2.8. Homologs of CD2 utilized in phylogenetic analysis
Species name

Protein

Gene Identifier #

Source

Arabidopsis thaliana

HDG7

AT5G52170

http://www.arabidopsis.org/

HDG1

AT3G61150

ANL2

AT4G00730

Solanum lycopersicum

CD2

GQ222185

http://solgenomics.net/

Solanum phureja

CD2
LOC Os09g35760

http://rice.plantbiology.msu.edu/

Oryza sativa

ROC6

LOC Os04g48070
LOC Os02g45250
Zea mays
Sorghum bicolor

OCL1

GRMZM2G026643

OCL3

GRMZM2G116658
Sb03g008090
Sb06g025750

http://www.gramene.org/
http://www.gramene.org/

Sb02g030470
Glycine max

Glyma07g08340

http://www.phytozome.net/

Glyma03g01860
Glyma09g40130
Glyma18g45970
Medicago truncatula
Vitis vinifera
Populus trichocarpa

Medtr7g083690

http://www.phytozome.net/

XP002272264

http://www.phytozome.net/

POPTR 0002s15610

http://www.phytozome.net/

POPTR 0014s07130
Ricinus communis

RC30174.t000366

http://www.phytozome.net/

Manihot esculenta

cassava4.1 001772m.g

http://www.phytozome.net/

cassava4.1 001764m.g
Malus domestica

HM122582

88

http://www.ncbi.nlm.nih.gov/

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CHAPTER III

KNOX genes influence a gradient of fruit chloroplast development through regulation of
GOLDEN2-LIKE expression in tomato

Satya Swathi Nadakuduti, William L. Holdsworth1, Chelsey L. Klein and Cornelius S. Barry2

(The Plant Journal in press)

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Abstract
The chlorophyll content of unripe fleshy fruits positively correlates to nutrient content
and flavor of ripe fruit. In tomato (Solanum lycopersicum) fruit, the uniform ripening (u) locus,
which encodes a GOLDEN 2-LIKE transcription factor (SlGLK2), influences a gradient of
chloroplast development that extends from the calyx end to the base of the fruit. With the
exception of the u locus, the factors that influence the formation of this developmental gradient
are unknown. In this study, characterization and positional cloning of the uniform gray-green
(ug) locus of tomato reveals a hereto unknown role for the class I KNOTTED1-LIKE
HOMEOBOX (KNOX) gene, TKN4, in specifying formation of this chloroplast gradient. The
involvement of KNOX in fruit chloroplast development was confirmed through characterization
of the Curl (Cu) mutant, a dominant gain-of-function mutation of TKN2, which displays ectopic
fruit chloroplast development that resembles SlGLK2 over-expression. TKN2 and TKN4 act
upstream of SlGLK2 and the related gene ARABIDOPSIS PSEUDO RESPONSE REGULATOR
2-LIKE (SlAPRR2-LIKE) to establish their latitudinal gradient of expression across developing
fruit that leads to a gradient of chloroplast development. Class I KNOX genes typically influence
plant morphology through maintenance of meristem activity but this study identifies a role for
TKN2 and TKN4 in specifically influencing chloroplast development in fruit but not leaves,
suggesting that this fundamental process is differentially regulated in these two organs.

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Introduction
In plants, many biochemical pathways, including those associated with photosynthesis
and synthesis of primary and specialized metabolites are located within the chloroplast. Leaves
are typically the primary source of photosynthate although it is estimated that photosynthesis by
fleshy fruits contributes up to 20% of the total fruit carbohydrate (Blanke and Lenz, 1989;
Hetherington et al., 1998). Enhanced chloroplast development and chlorophyll content of unripe
fruits increases their photosynthetic capacity leading to greater starch accumulation (Dinar and
Stevens, 1981; Powell et al., 2012; Sagar et al., 2013). Furthermore, the starch content of unripe
fruit is positively correlated with the sugar content in ripe fruit, which directly contributes to fruit
quality by influencing nutritive value and taste (Davies and Hobson, 1981). Thus, chloroplast
development and the carbohydrate status of unripe fruits impact the composition and quality of
ripe fruit.
In tomato, several loci influence fruit chloroplast development and the subsequent quality
of ripe fruit. Mutations at the high-pigment 1 and 2 (hp-1 and hp-2) loci, which encode negative
regulators of photomorphogenesis, possess increased chloroplast number and plastid
compartment size, leading to fruits with higher levels of chlorophyll and carotenoids (Mustilli et
al., 1999; Cookson et al., 2003; Liu et al., 2004; Kolotilin et al., 2007; Rohrmann et al., 2011).
Alteration of phytohormone biosynthesis and responses, including those associated with auxin,
ABA, cytokinins and brassinosteroids (BRs), can also positively impact chloroplast development
to influence ripe fruit quality (Martineau et al., 1994; Galpaz et al., 2008; Sagar et al., 2013; Liu
et al., 2014). In contrast, mutation at the uniform ripening (u) locus negatively impacts fruit
chloroplast development, due to loss of the green shoulder that surrounds the calyx end of
developing fruit, leading to ripe fruits with reduced sugar content (Powell et al., 2012). Positional

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cloning of u revealed a mutation in a tomato homolog of the GOLDEN 2-LIKE (GLK)
transcription factor of maize, designated SlGLK2 (Powell et al., 2012). GLKs are required for
chloroplast development (Fitter et al., 2002; Waters et al., 2008) and SlGLK2 is preferentially
expressed in the shoulder of the fruit, where in U/U genotypes, it is involved in enhancing
chloroplast development while ectopic expression of SlGLK2 results in homogeneously darkgreen fruit (Powell et al., 2012). Similarly, over-expression of ARABIDOPSIS PSEUDO
RESPONSE REGULATOR 2-LIKE (SlAPRR2-LIKE) in tomato, which is related to SlGLK2,
causes enhanced fruit chloroplast development (Pan et al., 2013). How these loci interact to
jointly influence fruit chloroplast development is not fully understood although recent evidence
suggests a role for both auxin and BRs in regulating GLK expression (Sagar et al., 2013; Liu et
al., 2014). Furthermore, except for the involvement of SlGLK2, the factors that establish the
latitudinal gradient of gene expression and chloroplast development in tomato fruit remain
undefined.
Fruits of the uniform gray-green (ug) mutant of tomato are similar to u and lack the green
shoulder surrounding the calyx (Bohn and Scott, 1945). In this study, positional cloning revealed
that UG encodes TKN4, a class I KNOTTED1-LIKE HOMEOBOX (KNOX) transcription
factor. KNOX genes typically influence plant morphology, and particularly leaf shape, through
maintenance of stem cell populations within apical meristems (Hay and Tsiantis, 2010).
However, this research identifies a previously undefined role for these homeobox proteins in
fleshy fruit development, and in particular in the establishment of a gradient of chloroplast
development through regulation of SlGLK2 expression.

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Results
Positional cloning of ug defines a role for KNOX in fruit chloroplast development
The ug mutant lacks the green shoulder surrounding the calyx end of developing fruit
(Figure 3.1A). Chlorophyll levels are reduced in ug fruit pericarp (Figure 3.1B-E). Individual
chloroplast development in ug fruit is also impaired with a reduction in size and thylakoids per
granum (Figure 3.1F-K). The phenotypes of ug are similar to fruits with the u mutant allele
(Powell et al., 2012) (Figure 3.1A-D, G and J). Genetic mapping assigned the ug locus to the
long arm of chromosome 1, with tight linkage to the marker C2_At1g56345. Fine mapping
refined the ug locus to a 14.6 kb interval between the markers Solyc01g100500 and
Solyc01g100520 that spans nucleotides SL2.40chr1: 82268632-82283258 and contains the single
gene, Solyc01g100510, which encodes the KNOX transcription factor, TKN4. Sequencing of
TKN4 from flower cDNA of UG/UG and ug/ug genotypes revealed a single T > A nucleotide
change within exon 2 of TKN4 corresponding to a phenylalanine to leucine substitution at
residue 137 (Figure 3.2A). Phenylalanine 137 represents an invariant amino acid within the
KNOX2 domain of diverse class I KNOX proteins (Figure 3.2B). The KNOX2 domain mediates
protein-protein interactions between homeodomain proteins (Bellaoui et al., 2001; Smith et al.,
2002). Genotyping of the SNP associated with the ug phenotype from LA4025 and Purple
Prince, which lack the green shoulder and are purported to carry the ug mutation implied that the
F137L substitution co-segregates with the mutant phenotype, indicating that the SNP is
conserved in ug/ug genotypes (Figure 3.2C and Figure 3.3).

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Figure 3.1. Fruit phenotype and chloroplast development in ug. A) Phenotypes of the u and
ug mutants compared to Ailsa Craig wild type (WT). The arrow highlights the presence of the
green shoulder in WT fruit. B) Chlorophyll content in the pericarp isolated from the shoulder (S)
and base (B) of WT, u, and ug fruit at 28 days post-anthesis (dpa). Data are presented as the
mean ± SEM of n=5 and means with different letters are significantly different (LS Means, p <
0.05). C-E) Confocal laser scanning microscopy images of WT, u, and ug fruit shoulder pericarp
showing chlorophyll autofluorescence (Scale bar = 10 µm). F-K) TEM of 28 dpa fruit
chloroplasts of WT, u, and ug (Scale bar for F-H = 2µm; I-K = 0.5µm).
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Attempts failed to genotype the ug mutant allele in the LA0021 accession, which lacks a green
shoulder but is reported to be a ug mutant (Figure 3.3). Similarly, amplification of TKN4 from
LA0021 flower cDNA was unsuccessful (Figure 3.2D). Subsequent amplification and
sequencing of TKN4 genomic fragments from LA0021 revealed the existence of a second mutant
allele that contains a 2.4kb internal deletion spanning nucleotides SL2.40chr1: 8228036582282785 of the tomato genome reference assembly (Figure 3.2E). Although mutations in Class
I KNOX genes typically cause phenotypes associated with altered morphology, and particularly
leaf shape (Parnis et al., 1997; Venglat et al., 2002; Hay and Tsiantis, 2010), the effect of ug is
subtle and plants carrying the ug mutant allele are indistinguishable from wild-type (WT) except
for the lack of the green shoulder in developing fruit.
No impact was observed on leaf shape or leaf chlorophyll in ug (Figure 3.4A-C). TKN4 is
expressed in flower pistils and mature green fruits but expression is low in leaves supporting the
lack of an altered phenotype in leaves of the ug mutant (The Tomato Genome Consortium, 2012)
(Figure 3.4D and E). As the ug and u fruit phenotypes are virtually identical (Figure 3.1A),
germplasm lacking a green shoulder was also genotyped for the u mutant allele. While LA4025
is homozygous for both u and ug mutant alleles, LA3539, LA0021 and the Purple Prince cultivar
possess the WT (U/U) allele (Figure 3.2C and Figure 3.3). These data indicate that the lack of a
green shoulder in these accessions is the result of mutations in TKN4 and not mutations at the u
locus.

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Figure 3.2. Genetic mapping of the ug locus. A) Genetic and physical map of the ug locus
identifying a single candidate gene, Solyc01g100510, which encodes TKN4. Numbers of
recombinant individuals between genetic markers are provided. Sequencing of TKN4 from wild
type (WT) (UG/UG) and mutant (ug/ug) plants identified a single nucleotide polymorphism that
converts phenylalanine 137 to leucine. B) Alignment of the KNOX2 domain of phylogenetically
distinct KNOX proteins. The invariant phenylalanine residue within this domain and its
substitution to leucine in ug/ug are shown in red. C) Correlation of L137 with the ug/ug
phenotype in selected accessions of tomato. D) RT-PCR amplification of TKN4 from Ailsa Craig
(WT) and LA0021 flower cDNA. E) TKN4 amplification from WT and LA0021 genomic DNA
highlighting 2.4kb deletion in LA0021. This deletion spans the first three exons of location of
TKN4 and is highlighted by the red bar in A).
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Figure 3.3. Genotyping of TKN4 and SlGLK2 in various tomato accessions. A) Fruit
phenotypes of the tomato accessions compared to Ailsa Craig wild type (WT). Note the
association of fruit shoulder with WT alleles of UG and U. Genotypes defined as UG/UG and
U/U contain the wild type alleles and possess a dark green shoulder whereas genotypes defined
as either ug/ug or u/u lack the shoulder. B) Genotyping tomato accessions using a cleaved
amplified polymorphic sequence marker digested by MseI. Genotypes defined as UG/UG show a
single 227 bp band whereas ug/ug genotypes show two bands corresponding to 173 bp and 54
bp. No amplification was observed from accession LA0021. C) Single base insertion causes
frameshift and a pre-mature stop codon, which results in truncation of SlGLK2 in u/u accessions.
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Figure 3.4. Leaf phenotypes and relative expression of TKN4. A) Leaf phenotypes of wild
type (WT), u, and ug. B) Leaf phenotype of WT and LA0021 in four-week-old plants. C)
Chlorophyll content in the leaf of WT, u, and ug (n = 5). D) Relative expression of TKN4 in
different plant organs (The Tomato Genome Consortium, 2012). E) Relative expression of
TKN4 in different organs of WT flowers. Means ± SEM with different letters are significantly
different (LS Means, p < 0.05).

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KNOX over-expression alters fruit development and positively impacts chlorophyll
accumulation
Additional support for a role for KNOX genes in regulating fruit chloroplast development
is evident from characterization of the Curl (Cu) mutant. Cu represents a dominant gain-offunction mutation resulting from ectopic expression of TKN2, leading to altered plant
morphology (Parnis et al., 1997). Developing fruit of Cu possess elevated chlorophyll levels that
extend across the entire surface of the fruit (Figure 3.5A-E). In addition to increased chloroplast
number and higher chlorophyll content in Cu fruit, the size of the chloroplasts is larger than in
WT and there is a general increase in the number of thylakoids per granum (Figure 3.5F-I). The
phenotypes of Cu fruit are reminiscent of SlGLK2 and SlAPRR2-LIKE over-expression lines
(Powell et al., 2012; Pan et al., 2013). To assess the potential relationship between Cu and the
SlGLKs and SlAPRR2-LIKE, the expression of these genes was examined in WT and Cu fruit
pericarp. The expression of TKN2, SlGLKs and SlAPRR2-LIKE is elevated in Cu fruit (Figure
3.5J-M). However, SlGLK1 expression, which is typically lower than SlGLK2 expression in
tomato fruit (Powell et al., 2012), was not statistically different between WT and Cu. These data
suggest that SlGLK2 and SlAPRR2-LIKE act downstream of KNOX activity to regulate fruit
chloroplast development.
In support of this hypothesis, fruit of a double mutant constructed from a cross between
Cu and u partially suppresses the Cu mutant phenotype (Figure 3.5N and O). Cu fruit are smaller
in size (WT=37.61g ± 2.53g; Cu=17.24g ± 1.69g, n=8) with a thinner pericarp and smaller cell
size that results in an increased density of cells per unit area (Figure 3.5D, 3P and Q). Congruent
with the phenotype of the ug allele (Figure 3.4A-C), the Cu allele does not impact leaf
chlorophyll content (Figure 3.6A) and while the expression of the SlGLKs remains unchanged,
the expression of SlAPRR2-LIKE is slightly elevated in Cu leaves (Figure 3.6B-E).
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Figure 3.5. Enhanced chloroplast development in Cu fruit. A and B) Variation in fruit
phenotype between Ailsa Craig wild type (WT) and Cu at 16 days post-anthesis (dpa). C and D)
Confocal laser scanning microscopy images of WT and Cu fruit shoulder pericarp showing
chlorophyll autofluorescence. (Scale bar = 10 µm). E) Chlorophyll content of WT and Cu fruit
pericarp (n=5). F-I) TEM of 28 dpa fruit chloroplasts of WT and Cu (Scale bar for F and G =
2µm; H and I = 0.5µm). J-M) qRT-PCR analysis of gene expression in WT and Cu fruit pericarp.
N) Fruit phenotype of Cu and Cu/u. O) Chlorophyll content of Cu and Cu/u fruit pericarp (n=3).
(P and Q) Pericarp thickness of WT and Cu fruit (n=6). Data are presented as the mean ± SEM.
Asterisks denote significant differences (***, p < 0.001; **, p < 0.01) as determined by
Student’s t tests. ns = not significant.

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Figure 3.6. Chlorophyll and relative expression of genes in Cu leaf. A) Chlorophyll content
of Ailsa Craig wild type (WT) and Cu leaves (n=5). B-E) Relative expression level of SlGLK2,
SlAPRR2-LIKE, SlGLK1 and TKN2 in WT and Cu leaves (n=3). Data are presented as the mean
± SEM. Asterisks denote significant differences (**, p < 0.01; *, p < 0.05) as determined by
Student’s t-tests. ns = not significant.

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KNOX loci establish gradients of SlGLK2 expression that impact fruit chlorophyll
accumulation
The presence of a green shoulder in immature tomato fruit highlights the existence of a
latitudinal gradient of chloroplast development that correlates with the increased expression of
SlGLK2 at the calyx end of the fruit relative to the stylar end (Powell et al., 2012). To assess the
role of KNOX genes in regulating the formation of this gene expression gradient and its
associated differential pattern of chlorophyll accumulation, fruit pericarp was divided into whole,
shoulder, middle and base, and the expression of SlGLK2, SlAPRR2-LIKE, TKN2 and TKN4
investigated (Figure 3.7A).
In WT fruit, chlorophyll levels are higher in the fruit shoulder when compared to the
middle and the base of the fruit (Figure 3.7B), and the expression of each gene matches the
gradient in chlorophyll accumulation with higher transcript levels observed in the shoulder
compared to the middle and the base, although this pattern is more pronounced for SlGLK2,
TKN2 and TKN4 (Figure 3.7C-F and Figure 3.8). The Cu and ug mutations disrupt both the
chlorophyll gradient (Figure 3.7B) and expression gradient that is present in WT fruit. For
example, the expression level of SlGLK2 is positively impacted by Cu and negatively by ug, and
the typical gradient of expression is disrupted (Figure 3.7C). In contrast, the ug locus does not
impact the expression of SlAPRR2-LIKE and while the expression of this gene is elevated in Cu,
the expression gradient is not disrupted (Figure 3.7D). Together, these data not only support the
role of KNOX genes in regulating both chloroplast development and gene expression gradients
but also highlights that while SlGLK2 and SlAPRR2-LIKE both positively impact fruit
chloroplast development (Powell et al., 2012; Pan et al., 2013), they are regulated by
overlapping, yet distinct mechanisms.

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Figure 3.7. Mutations at knox loci disrupt the gradient of SlGLK2 expression to impact
fruit chloroplast development. A) Depiction of the sampling strategy. B) Chlorophyll content
in the pericarp isolated from the whole (W), shoulder (S), middle (M) and base (B) of Ailsa
Craig wild-type (WT), Cu, and ug mutant fruit at 28 days post-anthesis (n=5). C-F) qRT-PCR
analysis of expression of SlGLK2, SlAPRR2-LIKE, TKN2 and TKN4 in the same tissue samples
as described in B) relative to the expression level in base of WT fruits (n=3). Data are presented
as the mean ± SEM and means with different letters are significantly different (LS Means, p <
0.05).
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Figure 3.8. Relative expression gradient of TKN2 within the fruit. qRT-PCR analysis of
expression of TKN2 in the pericarp isolated from the whole (W), shoulder (S), middle (M) and
base (B) of Ailsa Craig wild-type (WT), and ug mutant fruit at 28 days post-anthesis (n=3). Data
are presented as the mean ± SEM and means with different letters are significantly different (LS
Means, p < 0.05). This represents an alternative depiction of the data presented in Figure 3.7 to
highlight the gradient dependent expression of TKN2 in WT and ug fruit pericarp that are
masked by the elevated expression of TKN2 in the Cu mutant background.

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Several studies implicate phytohormones in mediating fruit chloroplast development in
tomato, and in the case of auxin and BRs, enhanced fruit chloroplast development is positively
correlated with increased SlGLK expression (Martineau et al., 1994; Galpaz et al., 2008; Sagar et
al., 2013; Liu et al., 2014). In addition, auxin gradients are involved in several developmental
processes and disruption of auxin signaling in the arf6/arf8 double mutant results in ectopic
KNOX expression and altered floral morphology (Benkova et al., 2003; Petersson et al., 2009;
Tabata et al., 2010). The potential role of auxin in regulating the KNOX-mediated fruit
chloroplast gradient in tomato fruit was investigated utilizing SlARF4-silenced lines, which
possess increased starch content, enhanced chloroplast development and elevated SlGLK1
expression (Sagar et al., 2013). The SlARF4-silenced lines were developed in the Micro-Tom
background (Kobayashi et al., 2014), which is homozygous for the u mutation and lacks a
functional SlGLK2. Unexpectedly, and unlike the u mutant allele in the Ailsa Craig background
(Figure 3.1A and B), a decreasing gradient of chlorophyll content from the shoulder to the base
of the fruit was observed in both Micro-Tom and an SlARF4-silenced line, with the latter
displaying the expected elevated chlorophyll content (Figure 3.9A). The expression of SlGLK2
and SlAPRR2-LIKE are elevated in the SlARF4-silenced line, particularly in the shoulder, and
whereas the expression of TKN2 and TKN4 was also enriched in the fruit shoulder, the difference
in expression levels between Micro-Tom and the SlARF4-silenced line are minor (Figure 3.9BE). These data highlight the existence of a gradient of chlorophyll accumulation in Micro-Tom
that occurs independently of SlGLK2 and suggests that SlARF4 negatively regulates SlGLK2 and
SlAPRR2-LIKE expression.

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Figure 3.9. Silencing of SlARF4 impacts fruit chlorophyll and expression of SlGLK2 and
SlAPRR2-LIKE. A) Chlorophyll content in whole (W), shoulder (S) and base (B) pericarp of
Micro-Tom and SlARF4 antisense lines at 21 days post-anthesis (n=5). B-E) Relative expression
level of SlGLK2, SlAPRR2-LIKE, TKN2 and TKN4 in same samples as described in A). Data are
presented relative to the expression level in base of Microtom fruits (n=3). Means ± SEM with
different letters are significantly different (LS Means, p < 0.05).
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The fruit phenotype of hp-1 is partially suppressed by u and ug
Fruits of the hp-1 mutant have increased chloroplast number, increased plastid
compartment size and a higher chlorophyll content (Cookson et al., 2003). The contribution of
KNOX and SlGLK2 to the hp-1 phenotype was assessed using a combination of single and
double mutants in both WT and the hp-1 background. Both u and ug partially suppress the hp-1
phenotype although suppression by u is greater than that achieved by ug and a slightly additive
phenotype is apparent when u and ug are combined (Figure 3.10A and B). Congruent with the
elevated levels of chlorophyll, the expression of SlGLK2, SlAPRR2-LIKE, TKN2 and TKN4 are
higher in hp-1 compared to WT (Figure 3.10C-F). Together, these data indicate that both
SlGLK2 and TKN4 contribute to the hp-1 mutant phenotype and are also suggestive of a role for
SlAPRR2-LIKE in determining the elevated chlorophyll content of hp-1 fruit, possibly via
regulation by TKN2. The reduced suppression of hp-1 by ug compared to u may be due to the
residual expression of SlGLK2 observed in the ug mutant (Figure 3.7C), which may indicate that
the ug allele (accession LA3416), which carries the F137L substitution, is not a complete null or
that additional unknown factors regulate SlGLK2 expression. Furthermore, evidence for the
involvement of SlAPRR2-LIKE or additional TFs in determining the elevated chlorophyll content
of hp-1 fruit is supported by the observation that while chlorophyll levels of the hp1/u mutant are
reduced compared with those observed in WT fruit, they are not reduced to the levels observed
in the u mutant (Figure 3.10B).

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Figure 3.10. The u and ug loci partially suppress hp-1. A) Variation in fruit phenotype
between single, double and triple mutant combinations of hp-1, u and ug at 21 days post-anthesis
(dpa). B) Chlorophyll content in the pericarp of genotypes in A) (n=5). C-F) Relative expression
level of SlGLK2, SlAPRR2-LIKE, TKN2, and TKN4 in wild type (WT) and hp-1 pericarp at 21
dpa. Data are presented as the mean ± SEM. Means with different letters are significantly
different (LS Means, p < 0.05) and asterisks denote significant differences (**, p < 0.01) as
determined by Student’s t-test.

117

Discussion
A role for KNOX genes in fleshy fruit development
KNOX genes influence multiple aspects of plant morphology and are typically expressed
within apical meristems where they maintain pluripotent cell populations required for organ
initiation (Jackson et al., 1994; Long et al., 1996; Byrne et al., 2002; Belles-Boix et al., 2006;
Hay and Tsiantis, 2010). Loss of KNOX activity can lead to inhibition of shoot formation,
whereas ectopic KNOX expression results in pleiotropic phenotypes including increased lobe
formation in simple leaves and increased complexity of compound leaves (Lincoln et al., 1994;
Hareven et al., 1996; Long et al., 1996). In addition to roles in shoot apical meristem formation
and leaf development, KNOX genes influence plant height, disrupt apical dominance and alter
patterns of cell division and elongation (Douglas et al., 2002; Venglat et al., 2002). The
Arabidopsis KNOX genes, SHOOT MERISTEMLESS (STM), BREVIPEDICELLUS (BP), KNAT2
and KNAT6 also influence inflorescence and carpel development together with architecture of
abscission zones (Ragni et al., 2008; Shi et al., 2011). The role of individual KNOX genes in
tomato remains obscure, but the identification of the underlying basis of the ug mutant has
revealed an atypical role for TKN4 in influencing fruit chloroplast development and specifically
the formation of the green shoulder (Figure 3.1). This role is supported through characterization
of the fruit phenotypes of Cu that result from ectopic expression of TKN2 (Figure 3.5).
TKN2 is closely related to the STM gene of Arabidopsis, which is required for both
vegetative and reproductive meristem development (Long et al., 1996; Scofield et al., 2007; Hay
and Tsiantis, 2010). As observed by TKN2 over-expression in the Cu mutant, ectopic expression
of STM also leads aberrant meristem activity and altered plant morphology (Parnis et al., 1997;
Gallois et al., 2002; Scofield et al., 2013). Within the shoot apical meristem, both TKN2 and STM
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function in part through modulation of hormone activity, and particularly through influencing the
levels of cytokinins and gibberellins (Hay et al., 2002; Jasinski et al., 2005; Yanai et al., 2005;
Hay and Tsiantis, 2010; Shani et al., 2010). At present the role of these hormones in contributing
to the altered phenotypes observed in the fruit of the Cu mutant (Figure 3.5), remain to be
defined. TKN4 does not possess an apparent ortholog in Arabidopsis, although previous
phylogenetic analysis suggested that it is closely related to FaKNOX1 from strawberry and
NTH1 of tobacco (Chatterjee et al., 2011). Manipulation of FaKNOX1 expression in transgenic
strawberry and Arabidopsis alters plant morphology, including leaf, flower and meristem
architecture (Chatterjee et al., 2011). However, altered expression of additional KNOX family
members was also apparent in FaKNOX1-silenced lines, raising the possibility that the observed
phenotypes could, in part, be due to altered expression of other strawberry KNOX genes
(Chatterjee et al., 2011). Similarly, over-expression of NTH1 in tobacco altered plant
morphology, manifest through crinkling of the leaves and altered phyllotaxy, phenotypes that are
typical of KNOX over-expression (Tamaoki et al., 1999). Interestingly, the ug mutant does not
display any obvious phenotypes related to organ morphology (Figure 3.4), which may be the
result of partial redundancy, a phenomenon often observed between KNOX genes (Byrne et al.,
2002; Belles-Boix et al., 2006) that is supported by the overlapping expression patterns of the
four tomato Class I KNOX genes TKN1-4 in different tissues and organs (The Tomato Genome
Consortium, 2012). However, functional specificity also exists between the tomato KNOX genes,
as although TKN2 and TKN4 display increased expression in the shoulder of the fruit (Figure 3.7
and Figure 3.8), TKN2 does not substitute for reduced TKN4 activity in the ug mutant.
Furthermore, characterization of Cu revealed a broader role for KNOX genes in fleshy fruit
development as TKN2 over-expression not only promoted chloroplast development but also

119

impacted fruit size, pericarp thickness and pericarp cell density (Figure 3.5). Identification of a
loss-of-function allele of TKN2 or fruit-specific gene silencing of this gene will facilitate
improved understanding of its role in both fruit development, including chloroplast development.
Differential regulation of chloroplast development in leaves and fruits of tomato
Both u and ug impact the gradient of chloroplast development in tomato fruit although
neither locus influences chloroplast development in leaves (Figure 3.1 and 3.4A-C). In the case
of the u locus, a lack of impact on leaf chloroplast development is likely due to redundancy and
the overlapping expression of SlGLK1 and SlGLK2 in tomato leaf (Powell et al., 2012) whereas
the lack of impact of ug on leaf chlorophyll content may be due to the absence of TKN4
expression in leaves (The Tomato Genome Consortium, 2012). However, over-expression of
TKN2 in the Cu mutant does not increase chlorophyll content in leaves but impacts chloroplast
development and chlorophyll accumulation in fruit (Figure 3.5 and Figure 3.6). These data
suggest differential regulation of chloroplast development in fruits and leaves of tomato, a
hypothesis supported by data indicating that over-expression of GLKs, and SlAPRR2-LIKE,
together with suppression of SlARF4, positively impacts fruit, but not leaf, chloroplast
development (Powell et al., 2012; Pan et al., 2013; Sagar et al., 2013). The underlying basis for
this differential regulation of chloroplast development in tomato is unknown but may be due to
saturation of chloroplast development in tomato leaves, as these contain approximately 20-fold
higher levels of chlorophyll than fruit (Figure 3.1B and Figure 3.4C) and therefore maybe less
amenable to further enrichment. In addition to the apparent differential regulation of chloroplast
development in leaves and fruits of tomato, parallel pathways regulate fruit chloroplast
development. For example, the impact of SlAPRR2-LIKE over-expression on fruit chloroplast
development is independent of SlGLK2, as experiments were performed in Micro-Tom, which is
120

homozygous for the u mutant allele (Pan et al., 2013; Kobayashi et al., 2014). Similarly,
although u and ug suppress the hp-1 mutant phenotype, they do not restore chlorophyll to levels
detected in the single mutants in the absence of the hp-1 (Figure 3.10A and B). This discrepancy
may be due in part to the elevated expression of SlAPRR2-LIKE in the hp-1 background (Figure
3.10D). Availability of SlAPRR2-LIKE mutant alleles or silencing of SlAPRR2-LIKE in WT and
mutant backgrounds will facilitate greater understanding of the role of this gene in chloroplast
development, together with its relationship to KNOX genes and SlGLK2.
TKN4 influences the gradient of chloroplast development in tomato fruit through
regulation of SlGLK2
Transcription factor gradients play important roles in the development of multicellular
organisms but remain poorly understood in plants (Stathopoulos and Levine, 2002; Sozzani et
al., 2010; Chen et al., 2012). In grasses, leaf differentiation proceeds basipetally and is
accompanied by a gradient of differential gene expression, chloroplast development and
photosynthetic competency (Leech et al., 1973; Li et al., 2010). Similarly, this study has
investigated a tissue-level macro-gradient of chloroplast formation that occurs during tomato
fruit development that leads to the formation of a green shoulder. The physiological role of this
chloroplast gradient is unknown but its formation is increased under shade conditions, and the
shoulder possesses higher photosynthetic capacity than the remaining pericarp tissues,
suggesting that this may be an adaptive trait that contributes to fruit photosynthesis (Smillie et
al., 1999). Manipulation of KNOX activity, either through mutation in TKN4 in ug or overexpression of TKN2 in Cu, disrupts the gradient of chloroplast development in tomato fruit by
altering the spatial expression of SlGLK2 and SlAPRR2-LIKE (Figure 3.7).

121

Figure 3.11. A proposed model for KNOX activity in developing tomato fruit. The KNOX
genes TKN2 and TKN4 positively influence SlGLK2 and SlAPRR2-LIKE expression to promote
fruit chloroplast development. Auxin, via SlARF4, represses the expression of SlGLK2 and
SlAPRR2-LIKE and brassinosteroids (BRs), via SlBZR1, promote SlGLK2 expression (Liu et al.,
2014). Data indicate that TKN2 influences the expression of both SlGLK2 and SlAPRR2-LIKE
whereas TKN4 only influences the expression of SlGLK2. HP-1 acts upstream of these genes to
repress their expression.

122

However, TKN2 and TKN4 possess overlapping and distinct activities and while overexpression of TKN2 influences the expression of both SlGLK2 and SlAPRR2-LIKE, reduced
TKN4 activity only disrupts SlGLK2 expression (Figure 3.7 and 6). These data promote a model
(Figure 3.11) suggesting the existence of distinct regulatory modules that influence chloroplast
development in tomato fruit and it will be of interest to determine whether TKN2 and TKN4
directly bind to regulatory regions of SlGLK2 and SlAPRR2-LIKE. Similarly, while suppression
of SlARF4 did not dramatically alter the expression of TKN2 and TKN4, both SlGLK2 and
SlAPRR2-LIKE expression was elevated (Figure 3.9). These data suggest multiple regulatory
inputs that influence the expression of SlGLK2 and SlAPRR2-LIKE, a hypothesis supported by
the recent report indicating that activation of BR responses leads to elevated expression of
SlGLK2 and enhanced chloroplast development (Liu et al., 2014). Although TKN4 influences the
latitudinal gradient of SlGLK2 expression in tomato fruit, TKN2 and TKN4 also display a
gradient-dependent expression pattern in developing fruit, suggesting that KNOX genes
themselves do not establish the gene expression gradient that exists in this tissue and that as yet
unidentified signaling pathways are required. Furthermore, KNOX proteins and their mRNAs are
also trafficked through plasmodesmata (Lucas et al., 1995), raising the possibility that the fruit
chloroplast gradient may, at least in part, also be regulated through cell-cell movement of these
proteins as well as their gradient-dependent transcript accumulation. The impact of gene
expression gradients in plants are not fully understood but this study has identified a role for
KNOX genes, whereby they regulate the gradient-dependent expression pattern of transcription
factors required for fruit chloroplast development.

123

Materials and Methods
Plant material, growth conditions and chlorophyll extraction
Seeds of tomato (Solanum lycopersicum) cultivars, mutants, and mutant combinations at
the uniform ripening (u), uniform gray-green (ug), Curl (Cu), and high-pigment 1(hp-1) loci
(Table 3.1), together with Solanum pimpinellifolium accession LA1589, were obtained from the
Tomato Genetics Resource Center, UC Davis or commercial vendors. Plants were grown in peatbased compost supplemented with fertilizer in greenhouses at Michigan State University, East
Lansing, MI under 16 h day (25°C) and 8 h night (20°C). Micro-Tom and the SlARF4-silenced
line (Sagar et al., 2013) were grown in a growth room under similar conditions. Flowers were
tagged at anthesis and fruits subsequently harvested at defined ages based on days post-anthesis.
Chlorophyll content was sampled from fruit pericarp and the terminal leaflet of expanding leaves
of 4 week-old plants as previously described (Nadakuduti et al., 2012).
Genetic mapping of the ug locus and genotyping
An inter-specific F2 population segregating for the ug locus was generated from a cross
between S. lycopersicum (ug/ug) (LA3539) and S. pimpinellifolium (UG/UG) (LA1589). Fruit of
60 F2 individuals were phenotyped for the presence or absence of a green shoulder and plants
were genotyped with genetic markers spanning the tomato genome. Subsequent fine mapping
was performed on further 1600 F2 individuals using chromosome 1 genetic markers (Table 3.2).
Genomic DNA was extracted as previously described (Barry et al., 2005) and details of genetic
and physical maps together with associated molecular markers are available through the Sol
Genomics Network. Genotyping tomato accessions (Table 3.1) for the ug mutant allele was
achieved using a cleaved amplified polymorphic sequence marker (Table 3.3) that was resolved

124

by digestion with MseI. Genotyping for the u mutant allele was achieved through sequencing of
PCR fragments, amplified using primers (Table 3.3) that span the single nucleotide insertion site
(Powell et al., 2012).
Molecular cloning and sequence analysis
Full-length cDNA clones of TKN4 were amplified and cloned from flower cDNA of wild
type and the ug mutant as previously described (Nadakuduti et al., 2012). Five independent
clones from each genotype were selected for sequencing. The entire TKN4 gene was sequenced
from LA0021 genomic DNA. Sequences from the region with unexpected band size containing
the deletion were cloned and four independent clones were sequenced and assembled using
Sequencher

version 4.7. Amino acid alignments were generated using MUSCLE (Edgar,

2004). Primers are listed in (Table 3.3) and additional sequences utilized in multiple sequence
alignments are provided in (Table 3.4).
qRT-PCR analysis
Expression analysis was performed according to (Nadakuduti et al., 2012) with the
exception that 10µL PCR reactions were assembled using a Biomek 3000 liquid handler and
amplified using an Applied Biosystems ABI Prism 7900HT Real-Time PCR System with the
following program: 2 min at 50°C and 10 min at 95°C, followed by 40 cycles of 15 s at 95°C and
1 min at 60°C. Primers are listed in (Table 3.3).
Microscopy
Chloroplast imaging using confocal laser-scanning microscopy (CLSM) and transmission
electron microscopy (TEM) was performed as previously described (Barry et al., 2012) with
minor modifications. CLSM images were captured using an objective with numerical aperture of
125

0.5 and maximum intensity projection images were generated from a confocal z-series acquired
through an average thickness of 170 µm in 10 µm increments. Samples for TEM imaging were
embedded in Spurr resin.
Statistical analyses
Statistical analyses were performed using SAS. The genotypic constituents were
evaluated by Student’s t-test and LS Means.

Acknowledgements
This research was supported by a Discretionary Funding Initiative award from Michigan State
University (MSU), a Strategic Partnership Grant from the MSU Foundation and by USDA
National Institute of Food and Agriculture, hatch project number MICL02265. W.L.H. was
supported by the MSU College of Agriculture and Natural Resources Undergraduate Research
Program, the Larry D. Fowler Endowment Undergraduate Research Stipend and an
Undergraduate Research Support Program Grant from MSU College of Natural Sciences. We
thank Prof. Mondher Bouzayen, University of Toulouse for providing seeds of the SlARF4
antisense line, Drs. Alicia Pastor and Melinda Frame of the Center for Advanced Microscopy at
MSU for their assistance and Dr. Rebecca Grumet for comments on the manuscript.

126

Table 3.1. Tomato accessions used in the study
Genetic background

Green
shoulder

Genotype

Source of material

LA2838A

Ailsa Craig

yes

U/U & UG/UG

www.tgrc.ucdavis.edu

LA0533

Codine Red

yes

U/U & UG/UG

www.tgrc.ucdavis.edu

LA0534

Lukullus

yes

U/U & UG/UG

www.tgrc.ucdavis.edu

LA0535

Rheinlands Rhum

yes

U/U & UG/UG

www.tgrc.ucdavis.edu

LA1589

S.pimpinellifolium

yes

U/U & UG/UG

www.tgrc.ucdavis.edu

LA0021

Uniform Globe

NO

U/U & ug/ug

www.tgrc.ucdavis.edu

LA3539

Ailsa Craig

NO

U/U & ug/ug

www.tgrc.ucdavis.edu

LA4025

Florida 7547

NO

u/u & ug/ug

www.tgrc.ucdavis.edu

Accession #

Purple Prince

NO

U/U & ug/ug

Gary Ibsen’s

LA3911

Micro-Tom

NO

u/u & UG/UG

www.tgrc.ucdavis.edu

LA3247

u (Ailsa Craig)

NO

LA3740

Cu (Ailsa Craig)

www.tgrc.ucdavis.edu

LA 3538

hp1 (Ailsa Craig)

www.tgrc.ucdavis.edu

LA3542

u/ug (Ailsa Craig)

www.tgrc.ucdavis.edu

LA3371

hp1/u (Ailsa Craig)

www.tgrc.ucdavis.edu

LA3416

hp1/ug (Ailsa Craig)

www.tgrc.ucdavis.edu

LA3593

hp1/u/ug (Ailsa Craig)

www.tgrc.ucdavis.edu

ARF4-AS1

(Sagar et al., 2013)

www.tgrc.ucdavis.edu

Sagar, M., Chervin, C., Mila, I., Hao, Y., Roustan, J.-P., Benichou, M., Gibon, Y., Biais, B.t., Maury, P., Latche,
A., Pech, J.-C., Bouzayen, M. and Zouine, M. (2013) Sl-ARF4, an auxin response factor involved in the control of
sugar metabolism during tomato fruit development. Plant Physiol., 161, 1362-1374.

127

Table 3.2. PCR based genetic markers flanking the ug locus
Gene / Marker
C2_At1g56345
TG197

Oligonucleotide Primer Sequence
Forward

5´-TCGGTGGCAACCACAGATTACCCAG-3´

Reverse

5´- ATCGAGTCTATTAGCTAAATGAAGCTC-3´

Forward

5´- GTACTACGAGAATTAAGGCATGGT-3´

Reverse

5´- GTGTAAGTTTCAAGGCATTCACGA-3´

MspI
SNP Marker C → T

Sequence 5´- GGTCTCATCTTGTTTCCTGTGGC-3´
Solyc1g100480
Solyc1g100500
Solyc1g100520
Solyc1g100530
Solyc1g100580
C2_At3g05810
C2_At5g02270
HBa57A01T7
SGN-U316057
T0276

Forward

5´- GTCTAGTAGTTATGAAGCTCCTGC -3´

Reverse

5´-GGATGAAAAGCAAGTTAGAATCT-3´

Forward

5´-GCCATTATCGAGCGAGTTTATTCC-3´

Reverse

5´-TCTGAACCCAATGATATGATCTCC-3´

Forward

5´-GGCTAAAGTAACTTTCTTTGTCC-3´

Reverse

5´-GGCATATGCATTTATGTATGCCTC-3´

Forward

5´-GTGAAATTATGTGAATATGCAGC-3´

Reverse

5´-TCAACAACCATGTCTAATAGGCTA -3´

Forward

5´-TGCACTGGTTCAAGAAATGGAGCA-3´

Reverse

5´-TAGCAACATGGCAAATGTAGAGC-3´

Forward

5´- GCCGCAGTTGTTTGAATTACAACC-3´

Reverse

5´- GCCCTTCACTTATGCATAGCACC-3´

Forward

5´- GGAGGAGGGAAGTTGCGTTTGC-3´

Reverse

5´- AACCTTAATTAGTTCGTCTCGTC-3´

Forward

5´-GTTAGATGATCAATATGGTAGTTTC-3´

Reverse

5´- GGTTTATTGAGGGATATATGCCGCC-3´

Forward

5´-GAGATGAATGAAATTGCTACTGC-3´

Reverse

5´-GGCGACTAGTGTGCCATATTATAGT-3´

Forward 5´-CTCGAGGAAGACGAGAACTTTGTC-3´

HpyCH4IV
HpyCH4V
Amplicon based indel
DraI
EcoNI
NheI
RsaI
RsaI
Amplicon based indel
AfeI

Sequences of the oligonucleotide primers utilized to amplify chromosome 1 genetic markers, together with the
appropriate restriction enzyme to resolve the marker between Solanum lycopersicum accession LA3539 (ug/ug) and
Solanum pimpinellifolium accession LA1589 (UG/UG). The TG197 marker is a SNP based marker resolved through
direct sequencing of the amplicons and Solyc1g100520 and SGN-U316057 are indel markers that were directly
resolved through gel electrophoresis of the amplicon.

128

Table 3.3. Oligonucleotide primers used in the study
Gene

Primer

Sequence

Use

TKN4

UG CAPS-F

5 -TCCAGCTCACGTCACAGTAGTGA-3

Genotyping

(Solyc01g100510)

UG CAPS-R

5 -GTTTCTTGAGTAACATAGAGTCA-3

Genotyping

TKN4-qF

5 -TATCGATGGCCTTATCCCACGGAA-3

qRT-PCR

TKN4-qR

5 -TCGATCCAGCACTTACACCTTCCA-3

qRT-PCR

UG inter-6F3

5'- CATGTGGGTCCGCCCTGTGATTC-3'

PCR

LA0021 deln-R2

5'- CATGCGCAGGCGTGCTTGTCAGC-3'

PCR

UG inter-6F1

5'-GACTTGGAACCAGTGGCGGATC-3'

Cloning

UG inter-9R

5'-GCCATCGATAGTGATTCTTCCACC-3'

Cloning

TKN4 ORF-F

5'-ATGATGGATGAATTGAGCAAAC -3'

RT-PCR

TKN4 ORF-STOPR

5'-GGAGGTACTGGAAATATTGGCACTTGA-3'

RT-PCR

SlGLK2

SlGLK2-F

5 -ATGCTTGCTCTATCTTCATCATTGA-3

Genotyping

(Solyc10g008160)

SlGLK2-R

5 -TTGAAGATGACTAGCAATGTTATGTCT-3

Genotyping

SlGLK2-qF

5 -CCTTACATGTTTGGGGGCATCCAC -3

qRT-PCR

SlGLK2-qR

5 -GGGGTGCAAATCAGAGGC -3

qRT-PCR

GAPDH

GAPDHQ-F

5 - ATGCTCCCATGTTTGTTGTGGGTG-3

qRT-PCR

(Solyc05g014470)

GAPDHQ-R

5 -TTAGCCAAAGGTGCAAGGCAGTTC-3

qRT-PCR

GAPDH-F

5'- GACAAGGCTGCTGCTCACTT-3'

RT-PCR

GAPDH-R

5'- GCTTGACCTGCTGTCACCAAC-3'

RT-PCR

TKN2

TKN2-qF

5 -CCATATCCATCGGAATCTCAG-3

qRT-PCR

(Solyc02g081120)

TKN2-qR

5 -TGGTTTCCAATGCCTCTTTC-3

qRT-PCR

SlGLK1-qF

5 -CCGTAAGCAGTGGTGATGAGTCTG-3

qRT-PCR

SlGLK1-qR
Solyc08g077230qF2
Solyc08g077230qR2

5 -AACCCGAACCTACATCCGAAGC-3

qRT-PCR

5'-GCAACTCTGCTGCTGAGATGAA-3'

qRT-PCR

5'-CGTTGCAGAACGAGTAGACTATGTAGT-3'

qRT-PCR

SlGLK1
(Solyc07g053630)
SlAPRR2-LIKE
(Solyc08g077230)

1.

2.
3.

4.

Ref

(1)

(2)
(2)
(3)

(4)
(2)

Balaji, V., Mayrose, M., Sherf, O., Jacob-Hirsch, J., Eichenlaub, R., Iraki, N., Manulis-Sasson, S.,
Rechavi, G., Barash, I. and Sessa, G. (2008) Tomato transcriptional changes in response to Clavibacter
michiganensis subsp. michiganensis reveal a role for ethylene in disease development. Plant Physiol., 146,
1797-1809.
Meir, S., Philosoph-Hadas, S., Sundaresan, S., Selvaraj, K.S.V., Burd, S., Ophir, R., Kochanek, B.,
Reid, M.S., Jiang, C.Z. and Lers, A. (2010) Microarray analysis of the abscission-related transcriptome in
the tomato flower abscission zone in response to auxin depletion. Plant Physiol., 154, 1929-1956.
Powell, A.L.T., Nguyen, C.V., Hill, T., Cheng, K.L., Figueroa-Balderas, R., Aktas, H., Ashrafi, H., Pons,
C., Fernandez-Munoz, R., Vicente, A., Lopez-Baltazar, J., Barry, C.S., Liu, Y.S., Chetelat, R., Granell,
A., Van Deynze, A., Giovannoni, J.J. and Bennett, A.B. (2012) Uniform ripening encodes a Golden 2-like
transcription factor regulating tomato fruit chloroplast development. Science, 336, 1711-1715.
Shani, E., Burko, Y., Ben-Yaakov, L., Berger, Y., Amsellem, Z., Goldshmidt, A., Sharon, E. and Ori, N.
(2009) Stage-specific regulation of Solanum lycopersicum leaf maturation by class 1 KNOTTED1-LIKE
HOMEOBOX proteins. Plant Cell, 21, 3078-3092.

129

Table 3.4. KNOX genes used for amino acid alignments
Species

Name

Gene identifier#

Source

Solanum lycopersicum

TKN4

Solyc01g100510

http://solgenomics.net/

Solanum lycopersicum

TKN3

Solyc05g005090

http://solgenomics.net/

Arabidopsis thaliana

STM

AT1G62360.1

http://www.arabidopsis.org/

Arabidopsis thaliana

KNAT2

AT1G70510.1

http://www.arabidopsis.org/

Arabidopsis thaliana

KNAT6

AT1G23380.2

http://www.arabidopsis.org/

Antirrhinum majus

invaginata

AY072736

http://www.uniprot.org

Fragaria vesca

FaKNOX

GU339211

https://www.ncbi.nlm.nih.gov/

Zea mays

ZmLG4

AF457121

https://www.ncbi.nlm.nih.gov/

130

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CHAPTER IV

Genetic mapping and morphological characterization of the tomato fruit developmental
mutants easy peel (ep) and peach (p)

138

Abstract
Tomato (Solanum lycopersicum) has long served as a model crop species to study fleshy
fruit development and tomato fruit mutants have significantly contributed to understanding the
molecular basis of fruit development and ripening. In this study, two monogenic tomato fruit
surface related mutants, peach (p) and easy peel (ep) were characterized and the mutant loci
mapped. Fruit of the p mutant is characterized by a dull epidermis with an uneven surface and
abundance of trichomes. Scanning electron microscopy (SEM) of the surface of p mutant fruit
revealed that the fruit surface contains ridges and the epidermal cells are small and conical,
unlike the wild type (WT) cells that possess an essentially smooth fruit surface. Analysis of
cuticle chemistry revealed increased epidermal wax abundance in p fruit cuticles compared to
WT. Genetic mapping positioned the p locus to within a 965 kbp region of chromosome 2. Fruit
of the ep mutant are identical to those of WT except that ripe fruits can be easily peeled incurring
less damage to the underlying sub-dermal cells. Anatomical characterization by SEM revealed
that the ep phenotype is probably due to modifications in cell layers just beneath the epidermis
rather than specific defects in the cuticle and epidermis. The ep locus was placed on chromosome
8 to within a 20 Mbp interval. Future identification of these mutant loci will facilitate greater
understanding of the underlying biological mechanisms that contribute to the morphology of the
fruit exocarp.

139

Introduction
Fruit ripening is a complex biological phenomenon that is extensively studied using
tomato (Solanum lycopersicum) as a model crop species. Several biochemical modifications
occur as a part of the fruit ripening process, including sugar accumulation, reduction in organic
acids, alteration in pigments responsible for development of color, production of volatiles
responsible for aroma profile, modifications to the ultra-structure and composition of cell wall
such as dissolution of the cell wall and middle lamella and water accumulation in the fruit,
leading to changes in texture and firmness (Brady, 1987; Liu et al., 2004; Saladie et al., 2007).
All these changes require extensive transcriptional and epigenetic reprogramming at the onset of
ripening (Alba et al., 2005; Osorio et al., 2011; Rohrmann et al., 2011; Zhong et al., 2013).
Water content in ripe fruit of tomato accounts for more than 90% of the total weight (Ho
et al., 1987) increasing the internal pressure, which is ultimately transmitted to the outer skin.
Therefore, the mechanical performance of the fruit skin is of considerable importance so that the
fruit can retain water and maintain physical integrity while undergoing these modifications. The
fruit skin is a protective layer comprised of the cuticle, epidermis and a variable number of
underlying sub-dermal cells, which together affect fruit appearance, postharvest handling and
storage (Bargel and Neinhuis, 2005; Lara et al., 2014). The mechanical properties of the fruit
skin are mainly attributed to the cuticle (Bargel and Neinhuis, 2005). Tomato fruit have a
prominent continuous cuticle that is approximately 10 µm thick and that lacks stomata (Vogg et
al., 2004). The tomato fruit cuticle is deposited on the exterior surface of the epidermal cells but
also surrounds their radial and tangential walls and can even extend beyond to the hypodermis
(Bargel and Neinhuis, 2005). The tomato fruit cuticle undergoes various changes in chemical
composition, thickness, elasticity and extensibility from anthesis to ripening (Baker et al., 1982;

140

Matas et al., 2004; Dominguez et al., 2011). The cuticle affects surface phenomena in fruits such
as the color, sheen, texture and uniformity. The surface of tomato fruits also contains epidermal
hairs known as trichomes. Type I and type VI glandular trichomes are predominantly present in
immature fruits and their density decreases as the fruits expand and the majority abscise prior to
maturity and ripening (Li et al., 2004; Mintz-Oron et al., 2008). Glandular trichomes synthesize
and secrete a variety of specialized metabolites, including terpenes, acylsugars and flavonoids
that are toxic or repellent to herbivores (Kennedy, 2003; Sallaud et al., 2009; Schilmiller et al.,
2009; McDowell et al., 2011; Weinhold and Baldwin, 2011). Recently, progress in fruit surface
biology, and especially the identification of enzymes and regulators involved in cuticle
biosynthesis, has been achieved through the characterization of mutants with altered cuticle
properties (Hovav et al., 2007; Leide et al., 2007; Isaacson et al., 2009; Nadakuduti et al., 2012;
Yeats et al., 2012; Shi et al., 2013; Yeats et al., 2014).
In this study, the phenotypes and map positions of the tomato fruit surface mutants easy
peel (ep) and peach (p) were investigated (Price and Drinkard, 1908; Lindstrom, 1925; Verkerk
and Contant, 1969). Fruits of the p mutant are dull and are covered by dense and unusual
persistence of trichomes even in ripe fruits. Phenotypic characterization of the fruit cuticles using
scanning electron microscopy (SEM) and biochemical analysis through Gas chromatography –
mass spectrometry (GC-MS) reveals that p has altered epidermal cell structure and cuticular
composition. Fruits of the ep mutant look identical to those of wild type Ailsa Craig (AC) plants
except that the ripe fruits can be peeled easily without much damage to underlying pericarp cells.
Inter-specific crosses of p and ep mutants generated with S. pimpinellifolium were performed to
produce F2 mapping populations that were screened with polymorphic markers to assign the
mutant loci to chromosomes 2 and 8, respectively. The aim of this research is to ultimately

141

isolate these mutant loci by positional cloning and perform functional analysis of each gene. As
the fruit surface is an important determinant of fruit quality the combined genetic and phenotypic
analyses will provide valuable insight into fruit surface biology.

142

Results
Phenotypic characterization of the peach mutant
At all stages of development, fruit of the peach (p) mutant are characterized by a dull
colored epidermis with an uneven surface that contains an abundance of trichomes (Lindstrom,
1925) (Figure 4.1A). Two morphologically distinct trichome types are typically evident on the
surface of tomato fruit, type I glandular trichomes, which have a long multicellular stalk with a
single celled gland at the tip and type VI trichomes which are shorter but are topped with a four
celled glandular head. The prevalence of both trichome types is increased in p mutant fruit. In
addition, the surface of p mutant fruits is uneven, the epidermal cells appear smaller in size, are
cone shaped and covered by an abundance of ridges, in contrast to WT fruits which possess a
fairly smooth fruit surface. Variation in the extracellular deposits on the ridged surface of p fruit
was also evident (Figure 4.1D-E). Although the fruits have an altered surface, cuticle thickness
appears normal except that the cuticle does not penetrate beyond the epidermal cell layer, as is
typically the case in WT fruit (Figure 4.1F-G). Furthermore, as a general observation, p mutant
fruit have a rubbery texture, and ripe fruits seldom crack, even when left to over-ripen on the
vine.

143

Figure 4.1. Fruit phenotype of peach (p) LA2357. A) Dull appearance of fruits of p. B-C) SEM
micrograph of mature green (MG) p fruit surface showing increased trichome density in p. D-E)
SEM micrograph of MG fruit cross sections of WT and p showing variation in the surface
morphology of epidermis. F-G) SEM micrograph of ripe fruit cross section of WT and p showing
extracellular cuticle (Scale bar in B-C = 1 mm, D-G= 20μm). Note the ridged appearance of the
epidermal surface in p fruit visible in G.
144

The genetic background of the peach mutant
At the initiation of this research, the genetic background of the p mutant was unknown,
which creates problems in making accurate phenotypic comparisons and drawing conclusions
from these comparisons. Therefore, a backcross strategy was initiated to introgress the p
mutation into the Ailsa Craig (AC) genetic background, a cultivar previously selected for
introgression of varied morphological mutants of tomato (Darby et al., 1978). The mutants were
selected for dullness and hairiness during the backcrossing and a BC2F2 population that contains
approximately 87.5% of the AC genetic background was derived. The BC2F2 p fruits are dull
throughout their development (Figure 4.2A-B) and have increased trichome abundance
compared to the WT (Figure 4.2C-D). They also retain the ridged and rough nature of their fruit
epidermal surface whereas the surface of WT fruit is smooth (Figure 4.2E-F). These
morphological differences in the fruit surface persist as the fruit ripens. Generally, as WT fruit
develops and ripens, trichomes desiccate and abscise but more trichomes are retained in ripe fruit
of the p mutant (Figure 4.3A-B). However, the uneven fruit surface observed in p mutant fruit
may, in part, be caused by the remnants of structures retained after the trichomes abscise,
particularly the base cells (Figure 4.3C-D). A prominent cuticle is also deposited on the surface
of ripe fruit of the p mutant, which is comparable to WT (Figure 4.3E-F).

145

Figure 4.2. peach (p) fruit has altered fruit surface. A-B) Mature green fruit and ripe fruit of
wild type (WT) and p from BC2F2 population respectively. Note the association of dullness in
both stages of fruit development. C-D) SEM micrograph of 16 days post anthesis (dpa) WT fruit
cross section and the same aged fruit of p showing many trichomes on the fruit surface
respectively. E-F) SEM micrograph of 16dpa fruit surface of WT fruit having a smooth surface
compared to ridged fruit surface in case of p fruit. (Scale bar in C-D = 100 μm, E-F= 10μm).

146

Figure 4.3. Variations in peach (p) ripe fruit surface. A-B) Ripe fruit surface of wild type
(WT) and p from BC2F2 population respectively. Note the retention of trichomes in p. C-D) SEM
micrographs of WT fruit surface compared to p respectively. Arrow highlights the bumpy fruit
surface in p. E-F) SEM micrograph of ripe fruit cross section of WT fruit having a smooth
surface compared to ridged appearance in case of p fruit. Arrow highlights the cuticle deposition
in p fruit. (Scale bar in A-B = 100μm, C-D = 10μm and E-F= 20μm).

147

Fruit of the peach mutant have altered cuticular wax composition
Based on the morphology of the p mutant, it is evident that it possesses several altered
phenotypes associated with the fruit surface (Figure 4.1 – 4.4). To investigate whether these
altered phenotypes are associated with differences in cuticular composition, wax and cutin load
of fruit cuticles of the BC2F2 lines of p and WT were analyzed at two developmental stages,
mature green stage (MG) and ripe stage (breaker + 10days).
Typically, tomato fruit wax consists of hydrocarbons, triterpenoids, fatty acids, alcohols,
sterols and coumarates (Baker et al., 1982; Vogg et al., 2004) and overall, the p fruit showed
considerably enhanced wax abundance compared to WT at both stages of fruit development.
This increase was manifest both through quantitative changes in components typically observed
in WT together with the appearance of a few minor components in p fruit that were not detected
in WT (Figure 4.4 and Table 4.1). The increased wax content in p is predominantly due to
elevated levels of alkanes and triterpenoids, the two major constituents of wax that together
contribute greater than 80% of the total wax load (Figure 4.4A-B, Table 4.1). Hydrocarbons
comprising of straight chain alkanes, branched chain alkanes and alkenes are major components
of wax which in MG fruit of p were 3.6 times and ripe fruit were 2.7 times higher than WT
(Table 4.1). Among the alkanes, nonacosane (C29) and hentriacontane (C31) are the major
alkanes at both stages of fruit development and these increased the most in p fruit. Dotriacontane
(C32) alkane, although not as abundant in WT, showed a proportionally greater increase than
C29 and C31 alkanes in p (Figure 4.4A). Triterpenoids, dominated by α, β and δ amyrins
increased nearly two-fold in both MG and ripe stages in p fruit (Figure 4.4B, Table 4.1). Alkanes
are generally associated with the epicuticular waxes and triterpenoids with intracuticular waxes

148

Figure 4.4. Cuticular wax constituents of fruits of WT and p. A) Hydrocarbon constituents of
fruit wax include straight chain alkanes, branched chain alkanes and alkene. B) Triterpenoid and
sterol constituents of fruit wax of WT and p. C) Total wax content at mature green (MG) and
ripe stages of fruit development in WT and p. Data represents mean values (n=5) ±SE. Asterisks
denote significant differences (*, p < 0.05; **, p < 0.01; ***, p < 0.001) as determined by
Student’s t tests.

149

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Table 4.1. Fruit wax composition of AC and p in μg dm
C chain
Wax class
Green fruit
length/compound
p
AC
Alkanes
C23:0
0.70 ± 0.001***
C25:0
1.29 ± 0.2***
C26:0
5.97 ± 0.01***

Iso alkanes

Anteiso alkanes
Alkenes
Alkanols

Alkenols
Fatty acids

Triterpenoids/
Sterols/
Coumarate

Ripe fruit
AC
0.75 ± 0.07
3.26 ± 0.9

p
0.97 ± 0.05
1.07 ± 0.09***
6.87 ± 3.0**

C28:0

2.24 ± 0.2

28.04 ± 5.7***

2.63 ± 0.5

26.35 ± 4.5***

C29:0
C30:0
C31:0
C32:0
C33:0
C30:0
C31:0
C32:0
C32:0
C33:2
C20:0
C22:0
C23:0
C24:0
C26:0
C27:0
C29:0
C32:0
C22:2
C24:2
C16:0
C18:0
C 20:0
C22:0
C24:0
trans - Coumarate
ψ Taraxasterol
Taraxasterol
delta amyrin
ß amyrin
α amyrin
Multiflorenol

65.21 ± 5.4
28.59 ± 1.4
136.97 ± 15.0
10.54 ± 2.7
6.76 ± 0.8
14.42 ± 5.6
1.70 ± 0.2
1.41 ± 0.1
1.95 ± 0.2
1.57 ± 0.3
0.52 ± 0.2
0.85 ± 0.1
3.23 ± 1.6
1.80 ± 0.8
21.76 ± 1.3
10.73 ± 2.4
17.66 ± 3.4
0.73 ± 0.2
33.35 ± 6.9
70.41 ± 3.3
37.29 ± 7.1
438.61. ± 50.8
242.43 ± 30.4
276.22 ± 53.6
88.07 ± 8.8

325.94 ± 38.8***
51.40 ± 4.0***
346.57 ± 33.3***
135.56 ± 53.2***
43.71 ± 8.6**
2.78 ± 0.5***
30.12 ± 2.3
1.85 ± 0.1
5.44 ± 0.7***
8.33 ± 2.6**
1.86 ± 0.02***
3.39 ± 0.5
0.98 ± 0.01
3.10 ± 0.7*
2.03 ± 0.3***
4.37 ± 0.9***
5.67 ± 0.6
11.54 ± 3.1***
1.36 ± 0.5
10.52 ± 3.1*
22.88 ± 9.8*
10.24 ± 4.6*
2.48 ± 0.9**
0.55 ± 0.04***
26.72 ± 3.2
61.02 ± 9.0
64.95 ± 10.1
830.88 ± 15.6**
405.76 ± 69.4*
452.22 ± 70.4**
79.14 ± 11.3

74.31 ± 4.9
40.52 ± 2.1
257.10 ± 9.9
40.82 ± 2.6
38.66 ± 2.7
1.39 ± 0.3
21.06 ± 1.1
3.34± 0.2
2.25 ± 0.0
30.72 ± 5.5
2.56 ± 0.7
4.79 ± 0.7
2.20 ± 0.5
2.23 ± 0.7
4.65 ± 0.5
5.41 ± 1.1
4.28 ± 0.5
8.25 ± 3.1
13.09 ± 2.4
4.75 ± 1.7
0.93 ± 0.5
20.39 ± 2.9
59.19 ± 1.9
54.13 ± 7.1
321.00 ± 63.4
317.44 ± 47.6
180.90 ± 48.2
71.01 ± 8.8

371.30 ± 52.4***
86.23 ± 8.7**
480.66 ± 60.2**
221.10 ± 43.0***
66.90 ± 10.2**
3.51 ± 0.6**
29.62 ± 2.8**
5.21 ± 1.5
10.44 ± 2.2**
92.27 ± 4.27**
1.83 ± 0.5
3.73 ± 0.3
3.82 ± 0.7
2.50 ± 0.7
1.74 ± 0.4***
5.64 ± 2.4***
14.25 ± 5.6*
14.84 ± 5.3*
4.40 ± 1.8
5.50 ± 2.5
18.76 ± 4.3*
7.91 ± 1.8
1.50 ± 0.3
0.87 ± 0.1***
2.60 ± 0.3***
24.14 ± 5.1
67.85 ± 12.2
72.94 ± 22.0
634.42 ± 36.3**
476.47 ± 53.3
482.31 ± 47.5*
84.85 ± 14.3

Data are shown as means ± SE (n = 5). Asterisks denote significant differences between genotypes of the same
developmental stage (*, p < 0.05; **, p < 0.01; ***, p < 0.001) as determined by Student’s t-tests.

150

(Vogg et al., 2004), illustrating a general increase in both types of waxes in p (Table 4.1). Two
other minor classes of wax components are fatty acids and fatty alcohols. There was an overall
increase in fatty acids but no major differences in fatty alcohols were evident between the two
genotypes (Table 4.1). Overall, the total amount of wax was doubled in both MG fruit (WT:p is
15.2:29.9 µg cm-2) and ripe fruit (WT:p is 15.9: 33.4 µg cm-2) in p compared to WT (Figure
4.4C).
In tomato fruits, cutin comprises 80% of the total cuticle weight, waxes account for
approximately 3% and the remainder is attributed to polysaccharides (Baker et al., 1982).
Tomato cutin is comprised of both C16 and C18 fatty acids, their oxygenated derivatives and cis
and trans coumaric acids, although nearly 70 - 75% of the total cutin is composed of 10, 16 di
hydroxy hexadecanoic acid (Table 4.2). Fruits of p at both MG and ripe developmental stages
exhibited a pattern of cutin deposition similar to WT, with the majority of the cutin distributed at
the MG stage. Limited variation in cutin composition is present between WT and p fruits and
these occurred in minor components. For example, 9-OH, 16-oxo hexadecanoic acid showed a
ripening-related increase from MG to ripe fruit in WT, whereas in p fruit it remained the same.
9, 18-diOH octadecanoic acid and C16:1 hexadecenoic acid levels were higher in p compared to
WT although these represent minor cutin monomers out of the total cutin load and their
contribution to the fruit phenotype is unclear. Overall, whereas the cutin fraction of the cuticle is
not dramatically altered in p compared to WT, the wax component is increased considerably,
translating to a reduced cutin monomer to wax ratio, of approximately 73:1 (961.5 : 13.25 µg cm2

) in ripe fruits of WT to 22:1 (835.6 : 38.60 µg cm-2) in p.

151

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Table 4.2. Cutin monomer composition of WT and p in μg cm (%)
Cutin monomer
cis - Coumarate

Green fruit
p
AC
3.5 ± 0.3 (0.5)
4.8 ± 0.6 (0.6)

AC
3.6 ± 0.3 (0.4)

Ripe fruit
p
5.2 ± 0.3 (0.6)**

trans - Coumarate

7.8 ± 0.7 (1.0)

10.9 ± 1.3 (1.7)

10.9 ± 1.0 (1.1)

12.1 ± 0.8 (1.5)

Hexadecanoic acid

0.8 ± 0.1 (0.1)

0.9 ± 0.1 (0.1)

1.1 ± 0.0 (0.1)

0.8 ± 0.1 (0.1)

Hexadecane 1,16 - dioic acid

6.0 ± 0.5 (0.8)

5.5 ± 0.5 (0.6)

8.3 ± 0.4 (0.9)

5.8 ± 0.5(0.7)*

C16:1 Hexadecenoic acid

1.6 ± 0.3 (0.2)

3.9 ± 0.4 (0.5)**

4.6 ± 0.2 (0.5)

3.2 ± 0.4 (0.4)*

16-OH Hexadecanoic acid

22.6 ± 2.2 (3.0)

21.7 ± 1.6 (2.4)

26.2 ± 1.5 (2.7)

21.9 ± 2.2 (2.6)

9-OH, 16-oxo Hexadecanoic acid

7.2 ± 1.0 (0.9)

12.4 ± 1.5 (1.4)***

21.5 ± 1.3 (2.2)

11.2 ± 1.9 (1.3)**

Octadecane 1,16 - dioic acid

1.7 ± 0.2 (0.2)

2.9 ± 0.2 (0.3)*

2.9 ± 0.2 (0.3)

2.9 ± 0.3 (0.3)

8-OH Hexadecane 1,16 - dioic acid

17.2 ± 2.1 (2.3)

20.7 ± 1.8 (2.4)

24.5 ± 1.2 (2.5)

22.3 ± 2.0(2.7)

18-OH Octadecanoic acid

2.9 ± 0.3 (0.4)

3.5 ± 0.2 (0.4)

3.5 ± 0.3 (0.4)

9, 18-diOH Octadecanoic acid

10.9 ± 1.5 (1.4)

4.2 ± 0.2 (0.5) *
20.7 ± 1.0
(2.3)***
625.0 ± 35.9
(71.7)
138.8 ± 11.9
(15.9)
871.6 ± 53.6

14.4 ± 0.9 (1.5)

17.8 ± 1.6 (2.1)

684.6 ± 31.7
(71.2)
160.1 ± 9.6
(16.7)
961.5 ± 41.9

576.8 ± 58.3
(68.7)
157.7 ± 14.1
(19.2)
835.6 ± 72.6

10, 16-diOH Hexadecanoic acid
Not identified
Total

568.4 ± 64.9
(75.1)
111.8 ± 18.3
(14.5)
759.0 ± 91.1

Data are shown as means ± SE (n = 5). Asterisks denote significant differences between genotypes of the same
developmental stage (*, p < 0.05; **, p < 0.01; ***, p < 0.001) as determined by Student’s t-tests

.

152

Genetic mapping of the peach locus
To initiate genetic mapping of the p locus, an inter-specific F2 mapping population
comprised of 128 individuals was developed from a cross between S.lycopersicum (p/p)
(LA2357) and S.pimpinellifolium (P/P). S. pimpinellifolium (LA1589) was chosen as the
mapping parent because of its close phylogenetic relationship with cultivated tomato, fertility of
the F2 individuals, which is essential for phenotyping a fruit-related trait, and availability of
polymorphic markers. As described above, p mutant fruits of accession LA2357 had a dull
surface, from mature green (MG) to ripe and denser persistent pubescence of fruit epidermis.
Dullness and hairiness segregated in the F2 population, where some dull fruits were not hairy and
some hairy fruits were not dull. The basis for this segregation remains unknown, but the dull fruit
phenotype was more consistent than the trichome phenotype and was therefore used for
phenotyping of the F2 mapping population. Previously, the p locus was assigned to chromosome
2 of the classical linkage map of tomato (Rick and Butler, 1956) and using a combination of
available (http://solgenomics.net/) and custom designed chromosome 2 genetic markers this
location was confirmed and the p locus was positioned between the LEOH113 (~85.0cM) and
LEOH174 (~120.0cM) markers on chromosome 2 (Figure 4.5). Additional markers within the
region were utilized to refine the p locus to an interval flanked by C2_At2g04700 (~97.0cM)
(SL2.40ch02:44259745..44264878) and T1480 (~100.6cM) (SL2.40ch02:45223579..45224990)
which, based on the tomato genome assembly (The Tomato Genome Consortium, 2012),
represents a 965 kilobase physical gap containing 139 predicted genes. The marker information
for this region is provided in Table 4.3.

153

Figure 4.5. Genetic linkage map of chromosome 2. Blue bar represents chromosome 2 with
genetic markers used in the study along with their centimorgan intervals on the left are shown.
The Red bar represents the region to which the locus is initially narrowed down. The numbers in
between adjacent markers represent the informative recombinant individuals from a total of 128
F2 plants. Further populating the region with markers led to narrow down the interval further,
highlighted by red bar.

154

Table 4.3. PCR based genetic markers used for mapping p locus
Gene / Marker
T1117
C2_At1g78690
LEOH 23.3
C2_At5g45950
SSR104
LEOH348
LEOH 113
C2_At4g36530
C2_At2g04700
cLED-19-B24
T1480
C2_At3g02220
SGN-U313660
LEOH174
C2_At5g14250
C2_At1g65720
C2_At3g28720
SGN-U575014

Oligonucleotide Primer Sequence
Forward 5'-CAACTCCGTC TTTGTCAGCC-3'
Reverse 5'-TGCTTTGTCG TCTGCACCCT-3'
Forward 5'-TCCAGAAGGGAAGGTCTGTCAAGAAG-3'
Reverse 5'-AGTCATGTACAGACATTTTTGTGCTGC-3'
Forward 5'-CTATGCGTTTGTCGGTCGT-3'
Reverse 5'-CAAGGTAGTTGAAGGTATGACCA-3'
Forward 5'-GCATCATTCTGGAAGCTGTCCA-3'
Reverse 5'-AGCTTTAAGAGCTTCATTAGCCA-3'
Forward 5'-TTCCATTTGAATTCCAACCC-3'
Reverse 5'-CCCACTGCACATCAACTGAC-3'
Forward 5'-TGTTTCCCTTCATTCATGCT-3'
Reverse 5'-CCAATTGGATAAATTGGTGGT-3'
Forward 5'-AAACAGAGGTGCCGAAGAAA-3'
Reverse 5'-GAGCTACAAGCAGCAAACCA-3'
Forward 5'-AGATTTGCTAGGCTTTGGTTGGAG-3'
Reverse 5'-AGTGAATCCCCAAACTGTCCTGC-3'
Forward 5'-ATGAGAACTCTTCAAGCTTCCACCTC-3'
Reverse 5'-TGACCACAGAAGTAACACCTTTGTCC-3'
Forward 5'-GGCACCATCAGAACTTCATGCAGA-3'
Reverse 5'-GTACCATTATGATAATGAGGTGTTA-3'
Forward 5'-ACCACCTTGGATGAATACCG-3'
Reverse 5'-TGCAACAGCTTTTCCCTCTC-3'
Forward 5'-TGTAGAGATCAAATTGATTGGAAGCG-3'
Reverse 5'-TATCCCGTTCTCGAGCATTCTTG-3'
Forward 5'-GTCCTAAGTCCATTTTCACATCAC-3'
Reverse 5'-GTCAAACAGAACATATCATCAAAGC-3'
Forward 5'-CGAGTCCGAGGAAGACTGAT-3'
Reverse 5'-TCAAGACAGACACGGATTGC-3'
Forward 5'-ATCTCATTCCCAGTTGCAAAGGAGC-3'
Reverse 5'-AGTCAGCATTGGAGCCACACCACG-3'
Forward 5'-CCTCGATTCCGTGCTTTCTTCGT-3'
Reverse 5'-GTCACGGATTGAGCTCGTCACCGA-3'
Forward 5'-ATCCATCCCCTGCTTTCACCAAG-3'
Reverse 5'-ACCATATCCTTAAAGGCCACAGACTG-3'
Forward 5'-GAATCTCCTCCAAGTTCCAGAGA-3'
Reverse 5'-GGCTTCATTATTTCTCCATCCAC-3'

Restriction
enzyme
BglII
AluI
Tsp509I
RsaI
indel
HpyCH4IV
NlaIII
Hpy188III
HpyCH4IV
MspI
AluI
HinfI
TaqI
indel
BamHI
HinfI
TfiI
HinfI

Sequences of the oligonucleotide primers utilized to amplify chromosome 2 genetic markers, together with the
appropriate restriction enzyme to resolve the marker between Solanum lycopersicum accession LA2357 (p/p) and
Solanum pimpinellifolium accession LA1589 (P/P). SSR104 and LEOH174 are indel markers that were directly
resolved through gel electrophoresis of the amplicon.

155

Phenotypic characterization of the easy peel mutant
Fruits of the easy peel (ep) mutant are indistinguishable from those of wild-type until the
onset of ripening. Then, as ripening proceeds, the skin of mutant fruits can be more easily
removed from the pericarp compared to that of wild-type and congruent with this easy-peel
phenotype, less of the underlying sub-dermal cells adhere to the skin (Figure 4.6). In addition, a
general observation made while growing this mutant is that the fruits are prone to cracking as
they ripen, which ultimately promotes opportunistic fungal infections. Although the cuticle is
often associated with fruit cracking (Chu and Thompson, 1972; Emmons and Scott, 1997;
Emmons and Scott, 1998; Matas et al., 2004; Bargel and Neinhuis, 2005; Dominguez et al.,
2011), there was no difference in fruit cuticle deposition in ep suggesting that the mutant
phenotype is due to an alteration in the properties of the epidermal or sub-dermal cells (Figure
4.6B-C). In support of this hypothesis, integrity of the fruit peel failed at the level of the subdermal cell layers of ripe ep fruit during preparation for SEM analysis (Figure 4.6D-E).
Genetic mapping of the easy peel locus
The ep mutant was originally recovered as a recessive allele in the progeny of thermal
neutron irradiated seeds of the cultivar Money Maker (Verkerk and Contant, 1969). To initiate
genetic mapping, an inter-specific F2 mapping population of 205 F2 individuals was developed
from a cross between S.lycopersicum (ep/ep) (LA3616) and S.pimpinellifolium (LA1589)
(EP/EP). Fruits of the F2 population were phenotyped for easy removal of fruit peel and clean
fruit peel. As the ep mutant was not assigned to a chromosome, a collection of markers that are
evenly spaced across the entire twelve chromosomes were used to genotype a subset of the F2
population that displayed the ep mutant phenotype at high confidence.

156

Figure 4.6. Fruit phenotypes of easy peel (ep). A) The ep mutant is readily peeled from the
underlying pericarp cells whereas pericarp cells adhere to the cuticle of wild type (WT) Ailsa
Craig (AC) fruit. B-C) Scanning electron micrographs (SEM) of ripe fruit cross sections of WT
and ep respectively. D-E) SEM micrographs showing the peel coming off of the ripe fruit cross
section of ep easily as compared to AC. Note the attachment of 2-3 layers of cells beneath the
cuticle as it peel off. (Scale bar in B-C = 2 mm, D-E= 20μm).

157

Linkage was observed between these high confidence mutant individuals and the
C2_At5g27390 marker, which is located on chromosome 8 (Figure 4.7). Following assignment
of the ep locus to chromosome 8, additional markers (Table 4.4) were mapped within the entire
population,

ultimately

placing

(SL2.40ch08:30094098..30099462)

the

locus
and

between

C2_At5g27390

cLER-5-P17

(20.5cM)
(51.75cM)

(SL2.40ch08:51752314..51754989) which represents a large physical distance of 21.66
megabases but only contains 465 predicted genes (The Tomato Genome Consortium, 2012).

Figure 4.7. Association of C2_At5g27390 marker to ep phenotype. Screening the mutant
phenotypes of F2 mapping population from a cross between S.lycopersicum (ep/ep) (LA3616)
with S.pimpinellifolium (P/P) with the maker C2_At5g27390 located on chromosome 8 shows a
strong correlation between the genotype and the phenotype, thus placing the EP locus on
chromosome 8. The 17 genotypes used are high confidence mutants from the F2 mapping
population.

158

Figure 4.8. Genetic linkage map of chromosome 8 showing the region of interest. Blue bar
represents chromosome 8 with genetic markers used in the study along with their centimorgan
intervals on the left are shown. The Red bar represents the region to which the locus is initially
narrowed down based on the informative recombinant individuals, represented by the numbers in
between adjacent markers from a total of 205 F2 plants. Further populating the region with
markers led to narrow down the interval further, highlighted by red bar.
159

Table 4.4. PCR based genetic markers used for mapping ep locus
Gene / Marker
TG176
SGN-U573854

Oligonucleotide Primer Sequence
Forward 5'-AGTAATAGCACTGCCCCACA-3'
Forward 5'-TTCGGCAAGTTTAGCCAAATA-3'
Forward 5'-TCCACAAATGTTGAAATTCATGCA-3'

Restriction
enzyme
DraI
RsaI

Reverse 5'-GAGAATCGGACGCAGATCTGAGATC-3'
SGN-U221657

Forward 5'-AGGTTTCAATGGTGGAGCAC-3'

MspI

Reverse 5'-ACAGCTGCTCGTTTCAGACC-3'
SGN-U229378

Forward 5'-AGGCAGTGGTTGATATACCTTTTGC-3'

NruI

Reverse 5'-TGTTCCCATATTCATACGGTTTCC-3'
C2_At1g30360

Forward 5'-GCTCTGTACGCGTAGTTTGTGTG-3'

HpyCH4V

Reverse 5'-AGGTGGTAGCCTGGTAGGTGAGG-3'
C2_At5g27390

Forward 5'-ATGGCCATGTCCACGCTCCTC-3'

RsaI

Reverse 5'-TGGGCTTAGCCTTATCTCCATATAG-3'
CT92

Forward 5'-GCCCTCTTACTATCTGTTGTTTG-3'

MspI

Reverse 5'-GACAAGTTGCTATACCAGGACCT-3'
C2_At4g33030

Forward 5'-CTCGGGACCATGGGGGAATATGG-3'

NsiI

Reverse 5'-AGTGGATGTCCAACAGCAGCCTG-3'
cLER-5-P17

Forward 5'-CCACATCCAGACTGAGTCACCAC-3'
Reverse 5'-CTCTTGGGCTTCGCTGAGTTGCT-3'

TaqI

cLED-6-M17

Forward 5'-GTTGCTCGTTTACGAGTACATGG-3'
Reverse 5'-CCTGTCACTAGCTCCAATAGCAC-3'

NruI

T1517

Forward 5'-CGGTAGTCCATACCTAAATAGCA-3'

smlI

C2_At5g20910

Reverse 5'-GCAAGGGCATTATTGTCTTAACG-3'
Forward 5'-GGCATATATTCCAGTCCTCAACC-3'
Reverse 5'-CCTTCTGCTTCACCATCTCTTCG-3'

Ase1

C2_At5g11450

Forward 5'-CCACTCTTGGCAAGCAATGGGAA-3'

RsaI

C2_At5g11490
TG513
C2_At2g24830
TG302

Reverse 5'-GCCCTTGCCCAGAGCATATTGGG-3'
Forward 5'-CAGTCCCGATCCTTCCACTGAAA-3'
Reverse 5'-CCTCCCATGCAAAACCGTCTTAC-3'
Forward 5'-CCTGGCTGGTTAGCTGGTCTGAT-3'
Reverse 5'-CCGGATGGACTCGAGGATCCAAG-3'
Forward 5'-CGTGGCATAGCTTCCAAGATGAT-3'
Reverse 5'-CATGCACTGCAAGCTGAGTATTG-3'
Forward 5'-CTCTCCGGGTGGCTATTACA-3'
Reverse 5'-TCTTGGGACTCCTCCTTTTCT-3'

RsaI
EcoRI
HinfI
AluI

Sequences of the oligonucleotide primers utilized to amplify chromosome 8 genetic markers, together with the
appropriate restriction enzyme to resolve the marker between Solanum lycopersicum accession LA3616 (ep/ep) and
Solanum pimpinellifolium accession LA1589 (EP/EP).

160

Discussion
A range of altered fruit surface properties contributes to the p mutant phenotype
Fruit of the p mutant display pleiotropic phenotypes pertaining to the fruit surface. For
example, there is an increase in the number of trichomes, altered cuticular wax composition and
a ridged fruit surface with smaller and conical shaped epidermal cells (Figure 4.1 - 4.4). Analysis
of the cuticle composition of the p mutant revealed that although cutin load did not change,
cuticular wax load increased. A recent analysis of ethyl methane sulfonate (EMS) generated
brightness mutants in tomato showed that all mutants that were recovered with a dull fruit
surface, displayed an increase in wax load compared to WT and interestingly had characteristic
morphological alterations in epidermal cells along with a rugged fruit surface, although no
correlation between cutin load and dullness was observed. The epidermal cells in these dull
mutants were smaller in size, less elongated and more conical shaped with higher cell density
than WT, which is equivalent to p fruit phenotypes (Petit et al., 2013). Noting that p fruit have an
abundance of trichomes, which are themselves surrounded by a cuticle layer (Mahlberg and Kim,
1991), there is a possibility that these could contribute to the increased wax phenotype. The cutin
load was not increased in the p mutant when compared to WT and it is possible that this may be
the result of the exhaustive delipidation phase employed during the cutin extraction process,
which may remove the trichomes. Light reflection from an object determines whether its surface
appears glossy/shiny or dull. In relation to the p mutant, surface wax as well as epidermal
trichomes are known to alter light reflection properties (Bargel et al., 2006). The difference in the
shape of the epidermal cells could also alter the light reflecting pattern of the fruit surface. With
a smooth surface, there is increased specular reflectance of incident light, just like reflection

161

from mirror, whereas light becomes diffused if the surface is bumpy and ridged, which could
make the fruit appear dull (Petit et al., 2013).
Previously characterized tomato fruit wax mutant, lecer6, defective in a β-ketoacylcoenzyme A synthase that is involved in wax biosynthesis and cultivar Delayed Fruit
Deterioration (DFD) demonstrated the importance of cuticular waxes, especially long chain
alkanes in the permeability of tomato fruit cuticles. The presence of higher amounts of alkanes
generally reduces cuticle permeability and limits the transpiration loss from fruit surfaces (Hovav
et al., 2007; Leide et al., 2007; Saladie et al., 2007). Interestingly, although the p mutant fruits
have high amounts of alkanes as well as total wax, the surface has more shriveled appearance
and the fruit texture is rubbery and not as juicy when ripe. Identifying the gene underlying the
mutant could possibly address questions pertaining to the relationship between epidermal cell
morphology and dull appearance of fruit together with the regulation of water relations in fruit.
An anatomical perspective of the easy peel phenotype
The ep fruits can be peeled without causing much damage to the underlying pericarp
cells. Based on the anatomy of the fruit pericarp cells, a study comparing ep genotypes with dark
green (dg) genotypes that are hard to peel, suggested anatomical differences between the
varieties. The fruits of ep genotypes have a steep cell size gradient near the outer surface and
enlarged intercellular spaces resulting in a noticeable separation layer along with thinning of
parenchyma cell walls in the outer pericarp (Chu and Thompson, 1972; Mohr, 1990) (Mohr,
1990).
Typically primary cell walls are composed of cellulose, hemicellulose and a glycan
polymer network of pectin, which is abundant in fruit cell walls (Brummell, 2006; Cosgrove and
Jarvis, 2012). Textural modification during fruit ripening mainly involves cell wall disassembly
162

of parenchyma cells, the major cell type in fruits, along with degradation of middle lamella that
cements adjoining cells. This is achieved by solubilization of cell wall components by cell wall
modifying enzymes including, polygalacturonase, pectin methyl esterase, pectate lyase and
expansins (Eliel and Rose, 2013). These alter the firmness, cell-cell adhesion and textural
properties of fruits as they ripen. In the ep mutant, the dissociation of peel could be due to a lack
of cell-cell adhesion, specifically in the outermost cell layers of the fruit pericarp. Identification
of the gene underlying ep mutant and its functional analysis will shed light on the regulation of
cellular processes in the outer fruit pericarp.
Noting that ep fruits almost always cracked when ripe suggests possible relationship
between cracking and easy peeling. The role of the cuticle in imparting resistance to fruit
cracking is well documented. The extent of cuticle deposition, high stiffness of the peel could
play a role in crack resistance (Chu and Thompson, 1972; Emmons and Scott, 1997; Emmons
and Scott, 1998; Matas et al., 2004; Bargel and Neinhuis, 2005; Dominguez et al., 2011).
However, anatomical analysis of the cuticle of ep fruit does not support its role as the underlying
basis of the mutant phenotype and instead suggests that disruption of the subdermal cells may
lead to the observed structural defects (Figure 4.6B-C).
Advancing genetic mapping strategies to identify the genes associated with ep and p loci
The genetic mapping experiments presented in this study were performed before the
release of the tomato genome sequence assembly. Although, both polymorphic markers and
some scaffold regions were available, there were regions, particularly those associated with the
ep locus, which could not be populated with molecular markers. The availability of the tomato
genome sequence should facilitate positional cloning efforts for these loci. In addition, next
generation sequencing, either in the form of genome re-sequencing or transcriptome sequencing
163

may be utilized, together with the current, or slightly refined, map positions to identify the
underlying genes. In particular, as the ep mutant was recovered following irradiation with
thermal neutrons, it is expected to be a deletion mutant. Furthermore, as the mutant phenotype is
only manifest during the ripening process, it is possible that candidate genes may be identified by
mining the mapping interval for genes whose expression changes during ripening or are
specifically expressed in fruit. Alternatively, comparative transcriptome sequencing of WT and
ep ripe fruit pericarp may identify chromosome 8 genes that are not expressed in the mutant due
to a possible deletion at the ep locus.
Problems associated with phenotyping of the F2 mapping populations were also
encountered during this study. In the case of the p locus, the F1 heterozygous plants from a cross
between S. lycopersicum (p/p) and S. pimpinellifolium (P/P) (LA1589) showed normal colored
fruits and the F2 showed a Mendelian phenotypic segregation of normal to dull fruits indicating
that dullness is completely recessive. However, the presence of trichomes did not co-segregate
with dullness, which initially made phenotyping a problem. This suggests a possibility of more
than one gene being associated with the p locus or perhaps the existence of genetic interactions
with loci derived from the S. pimpinellifolium mapping parent. For mapping the ep locus, the
segregating F2 plants from cross between S. lycopersicum (ep/ep) (LA3616) and S.
pimpinellifolium (EP/EP) (LA1589) showed variation in the degree of peeling that ranged from
easy to difficult. This suggests the possible existence of an intermediate phenotype between the
two homozygous parental phenotypes, which may occur if the ep allele is only partially recessive
to the WT allele or if the S. pimpinellifolium mapping parent alters fruit morphology of
individuals from within the F2 mapping population. In support of the latter hypothesis,
continuous variation in fruit phenotypes, which range from those observed in the S. lycopersicum
164

to those of the S. pimpinellifolium parents, are evident. Now that chromosomal regions pertaining
to the ep and p loci are identified, it may be possible to overcome ambiguous phenotypes
associated with interspecific F2 populations by developing new F2 mapping populations with the
appropriate S. pennellii introgression line (Eshed and Zamir, 1995). These lines carry
overlapping, marker defined chromosomal segments of S. pennellii that span the entire genome,
within a S. lycopersicum background. Thus, while polymorphisms and phenotypic variation can
be captured from the introgression of interest, utilizing these lines will create less phenotypic
variability within derived F2 populations.

165

Materials and Methods
Plant material and growth conditions
Seeds of the peach (p/p) (LA2357) and easy peel (ep/ep) (LA3616) mutants together with
Solanum pimpinellifolium (LA1589) were obtained from the Tomato Genetics Resource Center
(http://tgrc.ucdavis.edu/). Seeds of the wild-type Ailsa Craig cultivar were originally obtained
from the Glasshouse Crops Research Institute (Littlehampton, Sussex, UK). Plants were grown
in peat-based compost supplemented with fertilizer in greenhouses at Michigan State University,
East Lansing, MI under 16 h day (25°C) and 8 h night (20°C). Flowers were tagged at anthesis
and fruits subsequently harvested at defined ages based on days post-anthesis.
Genetic Mapping
Two inter-specific F2 populations segregating for the p and ep loci were generated from
crosses between S. lycopersicum (p/p) (LA2357) and S. pimpinellifolium (P/P) (LA1589) and S.
lycopersicum (ep/ep) (LA3616) and S. pimpinellifolium (EP/EP) (LA1589), respectively. Fruit of
128 and 205 F2 individuals were phenotyped for dull fruits and easy peeling, respectively and in
the case of the ep F2 population, plants were genotyped with genetic markers spanning the
tomato genome whereas genotyping was restricted to chromosome 2 genetic markers in the case
of the p F2 population. Genomic DNA was extracted from expanding leaves as previously
described (Barry et al., 2005). Details of genetic maps and molecular markers can be accessed
through the Sol Genomics Network (http://solgenomics.net). A combination of previously
defined cleaved amplified polymorphic sequence markers and custom markers with
experimentally determined polymorphisms between the S.pimpinellifolium and S. lycopersicum
parents of the F2 mapping populations were utilized for genetic mapping.

166

Extraction and Analysis of Cuticular Lipids
Extraction of tomato fruit surface waxes was accomplished through dipping fruit in
chloroform containing 1% methanol for 1 min. Internal standards of tetracosane, hexadecanol
and heptadecanoic acid (5 μg/mL each; Sigma-Aldrich, http://www.sigmaaldrich.com) were
added to these chloroform extracts and were then dried under nitrogen gas and derivatized using
BSTFA (N, O- bis-(trimethlysilyl) trifluoroacetamide) and pyridine at 110ºC for 10 min to
prepare trimethyl-silyl derivatives of wax components. After cooling, the wax derivatives were
dissolved in 1:1 heptane: toluene (1:1 v/v).
Compound identification was performed by GC-mass spectrometry (MS; EI mode) using
an Agilent Technologies 6850 GC, equipped with an HP-5 MS column connected to an Agilent
5975 mass spectrometer. Helium was used as carrier gas at 2 mL min−1 and oven temperature
was programmed from 120°C to 340°C at 10°C min−1. Splitless injection was used and ions were
collected in scan mode (40–800 atomic mass units) with peaks quantified on the basis of their
total ion current. The surface area of fruits was determined prior to wax extraction. The diameter
of the fruits were measured at three principal axes and fruit areas are calculated using formula for
an ellipsoid S= 4π ((ab)1.6 + (ac)1.6 + (bc)1.6 / 3)1/1.6 (Igathinathane and Chattopadhyay, 1998). The
fruit surface area measurements were validated in other experiments and were very comparable
with fruit areas measured using Tomato Analyzer version 2.1.0.0., (Brewer et al., 2006).
Following the chloroform dip, fruits were air dried and peeled. The fruit peel was scraped
to remove the underlying pericarp cells to obtain a clean cuticle. The fruit cuticles were
delipidated using chloroform: methanol (2:1 v/v) at room temperature for approximately 8-10
hrs. They were re-extracted with chloroform: methanol (1:2 v/v) overnight and finally with 100%
methanol for an hour. All the above extractions were performed by shaking and exchanging fresh

167

solvent periodically. Following exhaustive delipidation, the cuticles were air dried and once
completely dried, dry weight was recorded. The dried cuticles were ground to a fine powder
using liquid nitrogen and stored at -20°C until required.
Cutin composition was determined as previously described (Molina et al., 2006) using 50
mg of the finely ground delipidated cuticle powder. Methyl heptadecanoate and ωpentadecalactone (10 μg/mL each; Sigma-Aldrich, http://www.sigmaaldrich.com) were added as
internal standards. Base catalyzed depolymerization was carried out in methanol containing 15%
(v/v) methyl acetate and 6% (w/v) sodium methoxide by heat suspension at 60°C with periodic
vortexing for two hours. Glacial acetic acid, was added to adjust to pH 4.6, diluted brine solution
(0.5M sodium chloride) was used to wash the organic phase and then extracted with
dichloromethane. The organic phase was washed three times with brine and dried over sodium
sulfate. The product was dried under nitrogen gas and derivatized using acetic anhydride and
pyridine to corresponding acetyl derivatives. Cutin monomer identification was performed by
GC-MS as described above. Both cutin and wax compounds were quantified by gas
chromatography (GC)-flame ionization detector (6890N, Agilent Technologies) analysis of the
derivatized wax or fatty acid methyl ester products used a DB-5 capillary column. GC
temperature was programmed from 140°C to 310°C for cutin and to 330°C for wax, at 5°C
min−1. Samples were injected in split mode (330°C injector temperature). Quantification was
based on flame ionization detector ion current using peak areas relative to internal standard peak
areas. Both cutin and wax amounts were expressed on a per unit fruit surface area basis.

168

Scanning electron microscopy
To image fruit cuticles by scanning electron microscopy, fruits were hand-sectioned and
fixed for 30 min in 4% glutaraldehyde followed by 30 min incubation in 0.1 M phosphate buffer
(pH 7.4). The samples were dehydrated through a graded ethanol series and further processed
using a critical point dryer. Following desiccation, the samples were mounted onto aluminium
stubs and sputter coated with 30nm gold particles using a SC-500 sputter coater (Emscope
Laboratories, Ashford, U.K). The micrographs were captured using a 6400V scanning electron
microscope (JEOL, Tokyo, Japan).
Statistical analyses
Statistical analyses were performed using SAS (SAS Institute, www.sas.com). The
genotypic constituents were evaluated by either Student’s t-tests or Least Squares Means
analysis.

169

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170

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CHAPTER V

Conclusions and perspectives

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One of the main challenges facing biology in this post genomic era is to assign molecular
functions to the numerous uncharacterized genes discovered as a result of genome sequencing
projects. This holds true for the various fruit genomes sequenced, including tomato, where
functional analysis of the majority of the encoded genes is lacking. Several approaches to predict
or determine unknown gene function can be utilized, for example gene-to-gene co-expression
analyses, predictions of sub-cellular localization and large scale determination of protein-protein
interactions each provide correlative analysis of potential gene functions. However, in order to
understand the function of specific genes, gene knock-out or knock-down approaches are
required and in many fleshy fruit bearing species, these approaches remain slow and laborious.
Forward genetics-based approaches have the advantage of beginning with a mutant of biological
interest but the disadvantage that the underlying gene is unknown. However, in the case of
tomato, the combination of high-density genetic maps and a reference genome sequence,
facilitates the process of identifying genes associated with mutant phenotypes. The studies
presented in this dissertation focus on the characterization of four monogenic mutants of tomato
that alter different aspects of fruit development. These mutants were characterized using a
combination of physiological, molecular and chemical analyses, they were mapped to the tomato
genome and in two cases the underlying genes were identified and characterized.
CD2, a master regulator of epidermal cell function
The sticky peel (pe) mutant of tomato disrupts several epidermis-related phenotypes,
including fruit and leaf cuticle biosynthesis, anthocyanin formation, stomatal density and
trichome formation (Nadakuduti et al., 2012). Map based cloning identified pe as a new mutant
allele of CUTIN DEFICIENT2, a class IV homeodomain-leucine zipper (HD-ZIP IV) family
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transcription factor (TF). Previously, CD2 was shown to be required for tomato fruit cuticle
formation (Isaacson et al., 2009) but this research revealed a broader role for this TF in plant
development. Furthermore, characterization of the anthocyaninless 2 mutant of Arabidopsis
(Kubo et al., 1999), which encodes a CD2 homolog, revealed that this mutant also exhibits a
mild perturbation in cuticle biosynthesis and defined a regulatory link between cuticle and
flavonoid biosynthetic pathways operating in epidermal cells.
The Class IV homeodomain-Leu zipper (HD-ZIP IV) family of TFs is unique to the plant
kingdom and is found from moss to higher plants but is not present in algal genomes suggesting
that the presence of these genes pertains to the evolution of terrestrial plants (Mukherjee et al.,
2009; Javelle et al., 2011). This hypothesis is supported by the epidermal-specific expression of
the majority of the characterized family members and through functional characterization of a
limited number of family members that all disrupt epidermal-cell related development and
metabolism (Valle et al., 1997; Dong et al., 1999; Ingram et al., 2000; Ito et al., 2002;
Swigonova et al., 2004; Nakamura et al., 2006; Guan et al., 2008; Isaacson et al., 2009; Javelle et
al., 2011; Chew et al., 2013). No comprehensive analysis of the HD-ZIP IV family has thus far
been performed in tomato and the only characterized HD-ZIP IV genes are CD2 and Woolly
(Wo), a spontaneous mutation that promotes trichome formation and causes embryo lethality in
its homozygous state (Yang et al., 2011). There are 37 predicted HD-ZIP IV genes present in the
tomato genome, some of which are present as tandem duplications within the genome and
functional characterization of these genes would represent a major effort but would provide
insight into their role in plant development and epidermal cell function.
In addition, identifying the target genes of CD2 is critical to elucidate the transcriptional
regulatory network in which it participates. Genome wide identification of target genes can be
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achieved using chromatin immunoprecipitation (ChIP) coupled with deep sequencing (ChIP-seq)
using a purified antibody against CD2 or by an epitope-tagged version of CD2. Studies involving
genome-wide characterization of the HD-ZIP IV family in diverse species revealed that these
TFs are associated with epidermis-related expression and/or function (Javelle et al., 2011) and QRT-PCR analysis in tomato, performed on stem peel, core and whole stem indicates that CD2 is
preferentially expressed in stem epidermis (Nadakuduti et al., 2012). Therefore to perform the
above experiments, the peel tissue could be separated from fruit or stem of WT and pe mutant
samples to increase specificity. From the ChIP-seq data it should be possible to identify the in
vivo binding sites of CD2 near the candidate genes and this approach could be coupled with
transcriptome sequencing of WT, mutant and CD2 over-expression lines to reveal direct and
indirect targets of CD2. In particular, placing the function of CD2 into context with SlSHINE3 as
well as homologs of regulators of cuticle biosynthesis yet to be identified in tomato, including
members of other SHN TFs, MYB TFs that are biotic and abiotic stress responsive along with
HDG1 in Arabidopsis, and WXP1 identified in Medicago sativa (Zhang et al., 2005; Raffaele et
al., 2008; Seo et al., 2009; Seo and Park, 2010; Oshima et al., 2013) will facilitate identification
of distinct and overlapping regulons that influence epidermal cell function and in particular the
biosynthesis of cuticular lipids.
Uncovering the genetic basis of fruit chlorophyll gradients
Many fruits develop as green photosynthetic organs before they begin to ripen and
exhibit gradients of chlorophyll accumulation that are visibly obvious. These chlorophyll
patterns on fruits are intriguing and specific to each fruit, however limited knowledge exists on
the genetic mechanisms that control chloroplast development and chlorophyll accumulation
patterns in developing fruits. In tomato, a latitudinal gradient of chloroplast development exists,
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forming a green shoulder at the peduncle end of the fruit. Although, there has been a tremendous
advancement in regulation of fruit ripening, there is a knowledge gap that is particularly evident
for transcription factors acting during the early stages of fruit development. Chloroplast
development and fruit photosynthesis are crucial in determining the sugar, nutritional
composition and antioxidant capacity of the ripe fruit (Dinar and Stevens, 1981; Powell et al.,
2012; Sagar et al., 2013). Hence the fruit shoulder has a positive impact on the ripe fruit nutrient
content (Powell et al., 2012). One of the components of the regulatory network for fruit
chloroplast development was recently identified as the GOLDEN 2-LIKE transcription factor,
SlGLK2 (Powell et al., 2012).
Our research identified the gene that underlies the uniform gray-green (ug) locus of
tomato, which possesses a mutant phenotype that is virtually identical to that of the u locus. UG
encodes a previously uncharacterized class I KNOTTED-LIKE homeobox (KNOX) gene, TKN4,
providing the first evidence for involvement of KNOX genes in fleshy fruit development and in
particular, chloroplast development. The functional relationship between KNOX and fruit
chloroplast development was confirmed through characterization of the Curl (Cu) mutant, a
dominant gain-of-function mutation in the KNOX gene, TKN2 (Parnis et al., 1997), which
displays ectopic fruit chloroplast development. Furthermore, chlorophyll levels were not
influenced in the leaves of the ug or Cu mutants suggesting that separate pathways influence
chloroplast development in leaves and fruit of tomato. This hypothesis is supported by functional
characterization of SlAPRR2-LIKE and SlARF4, which also influenced fruit chloroplast
development but not leaf chloroplast development (Pan et al., 2013; Sagar et al., 2013). Gene
expression analysis indicates that TKN2 and TKN4 act upstream of SlGLK2 and SlAPRR2-LIKE
and influences their gradient-dependent expression, whereby transcript abundance is higher in

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the calyx end of the fruit and this stimulates shoulder formation. Recently, RNAseq analysis of
WT fruits at different stages of development revealed that latitudinal gene expression gradients
are robust and extend beyond the TF regulon identified in this study to encompass additional
classes of genes (Nguyen et al., 2014). Current understanding of the role of KNOX genes in
fleshy fruit development, and particularly chloroplast development is in its infancy and there are
several different approaches that could be pursued to increase the understanding of these TFs
with the ultimate aim of defining the regulatory framework in which they function.
To acquire further insight into establishment of chloroplast gradients and the specific role
played by KNOX genes in this process, generating transcriptome data from green fruit of Cu, ug
and WT by RNA-seq from tissue collected from the shoulder, middle and base portions of the
fruit could be pursued. We anticipate that the gradient of gene expression that correlates with
chloroplast development will be disrupted by ug and constitutively activated in Cu as a result of
reduced and enhanced KNOX activity, respectively. In addition, this dataset may provide insight
into potential broader roles of KNOX genes in fleshy fruit development and will therefore aid in
further expanding the current model of the KNOX-mediated regulatory network in tomato fruit
development.
SlGLK2 and SlAPRR2-LIKE may represent direct targets of KNOX and in the case of
SlAPRR2-LIKE, there is a repeated TGAC motif within the promoter sequence. Although KNOX
binding sites are often degenerate, genes with duplicated TGAC motifs within their regulatory
sequences often represent KNOX targets (Bolduc and Hake, 2009; Bolduc et al., 2012; Lin et al.,
2013). The ability of KNOX proteins to bind to regulatory regions of SlGLK2 and SlAPRR2LIKE can therefore be tested using a combination of chromatin immunoprecipitation (ChIP),
electrophoretic mobility shift assays (EMSAs) and transient expression assays. It is also possible

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that KNOX-mediated regulation of SlGLK2 and SlAPRR2-LIKE expression occurs through an
indirect mechanism. For example, ChIP-seq analysis of maize KNOTTED-1 revealed that it
targets hundreds of loci (Bolduc et al., 2012). Similarly, a ChIP-seq approach will be useful for
characterizing potentially broader roles for both TKN2 and TKN4 in fruit development and may
be necessary to gain further insight into this TF regulatory network if their impact on SlGLK2
and SlAPRR2-LIKE expression occurs through an indirect mechanism.
To date, the functional analysis of KNOX genes in fleshy fruit development has focused
on available mutant lines and has been restricted to reduced function of TKN4 and a gain of
function of TKN2. It would also be useful to examine the reciprocal activities. For example,
silencing of TKN2 may provide insight into the broader role of this gene in fruit development and
over-expression of TKN4 would allow the assessment of overlap with TKN2 and in particular to
determine whether over-expression of TKN4 mimics the Cu mutant phenotype. Manipulation of
KNOX activity can lead to highly pleiotropic phenotypes associated with altered plant
morphology as is observed in the Cu mutant. Therefore, fruit-specific manipulation of both
TKN2 and TKN4 may overcome these pleiotropic phenotypes, while focusing specifically on
those related to fruit development. A series of vectors for both fruit-specific silencing and overexpression are available that could facilitate such an analysis (Fernandez et al., 2009). In
addition, KNOX genes can act redundantly (Jackson et al., 1994; Long et al., 1996; Parnis et al.,
1997; Vollbrecht et al., 2000; Byrne et al., 2002; Belles-Boix et al., 2006). To counteract this
phenomenon, Shani et al., 2009 fused TKN2 to the EAR repressor motif of the Arabidopsis
SUPERMAN gene (Hiratsu et al., 2003; Shani et al., 2009). The EAR motif acts as dominant
repressor by suppressing the expression of target genes even in the presence of redundant
transcription factors, thus resulting in a dominant loss of function phenotype. This approach was

182

utilized for examining early leaf development and shape in tomato (Shani et al., 2009) but could
also be modified to encompass fruit-specific promoters and the resulting transgenic fruit
evaluated for altered development and potential changes to gene expression gradients.
KNOX proteins function by interacting with BEL1-Like (BELL) homeodomain proteins
and form heterodimers to target several genes by directly binding to the promoters of those genes
(Bellaoui et al., 2001; Smith et al., 2002; Smith and Hake, 2003). These interactions occur
through the MEINOX and SKY/BELL domains of KNOX and BELL proteins, respectively. A
single KNOX protein can interact with multiple BELL proteins, potentially providing flexibility
and specificity to signal outputs (Bellaoui et al., 2001; Smith et al., 2002; Hackbusch et al.,
2005). The role of BELL genes in fleshy fruit development is currently unknown and mining of
tomato transcriptome data revealed that 12 BELL genes are expressed at different stages of fruit
development (The Tomato Genome Consortium, 2012). This obviously creates the potential for
genetic redundancy and coupled with the potential complexity of interactions between KNOX
and BELL proteins, it will be difficult to determine the individual roles of these proteins in fruit
development. In the absence of defined mutants at these loci or the resources to attempt to
silence each gene singly, or in combination, it may be possible to also utilize an EAR domain
fusion to assess the potential impact of these genes on fruit development and gene expression
gradients.
Identification of loci underlying ep and p mutants will contribute to better understanding
of fruit exocarp
The peach (p) and easy peel (ep) mutants are the fruit developmental mutants of focus in
chapter IV. Fruits of the p mutant are dull and covered with dense trichomes, which persisted
even in ripe fruits. The shape and size of the epidermal cells are altered in p mutant fruits and
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they also display altered cuticular composition. This implies that the p mutant has pleiotropic
phenotypes pertaining to the fruit epidermis. Genetic mapping to isolate both loci is in progress.
We encountered problems in phenotyping p mutant, as fruit dullness and presence of trichomes
were segregating in the interspecific F2 mapping population. As the map position for this locus is
narrowed down to 1 Mbp on chromosome 2, developing new F2 mapping populations with the
appropriate S. pennellii introgression line will reduce this ambiguity of phenotyping and allow us
to fine map the locus fairly quickly.
Fruits of ep mutant can be peeled with limited damage to the underlying pericarp cells.
The dissociation of the fruit peel could be due to a lack of cell-cell adhesion, specifically in the
outer cell layers of the fruit pericarp. SEM micrographs suggest that the phenotype is beneath the
epidermis probably in parenchyma cells as fruit peel includes epidermis and some layers of
collenchyma cells. Genetic mapping placed the locus on chromosome 8. For further narrowing
down the interval, we have high confidence mutants in the F2 mapping population which will aid
in using the same population. With the availability of the reference genome of tomato and the
draft genome of S. pimpinellifolium, one of the parents of our F2 mapping population, it should
be possible to design and add more molecular markers to the mapping interval and increase the
mapping population to refine the map position. Comparative transcriptome sequencing of WT
and ep could also be utilized to detect the missing transcript in the mutant, due to a possible
deletion at the ep locus. The incorporation of laser capture micro dissection into this
experimental strategy could facilitate this approach as it would allow the precise isolation of the
exocarp tissue along with layers of parenchyma cells where we observe the peel dissociation to
occur. This method of sample collection for RNA-seq experiments has already been utilized to
obtain cell-specific transcripts in tomato fruit pericarp at an early stage of fruit development
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(Matas et al., 2011). However, some optimization may be required to fix and section tissues at a
later stage of fruit development when the ep phenotype is observed due to increased cell
expansion and ripening-induced cell wall disassembly. Identifying both p and ep loci will
enhance understanding of the molecular basis of fruit exocarp development and in the case of ep
may yield additional insight into the metabolic changes that occur during fruit ripening.
Importance of gene discovery in fruit crops and its impact on science and agriculture
Fruits form an essential component of the human diet as they provide minerals, vitamins,
antioxidants, carbohydrates and fiber to enrich the nutrient content along with enhancing our
overall enjoyment of food through the sensory pleasures of sweetness, color, flavor and aroma.
Tomato is a nutritious fruit commonly used as a vegetable crop that is grown throughout the
world. Tomato fruits are specifically rich in vitamins A, C and folic acid and contain a wide
array of carotenoids such as lycopene, beta-carotene, gamma-carotene, phytoene as well as
several minor carotenoids that possess antioxidant properties (Beecher, 1998). Additionally,
tomato serves as an excellent model system for studying fleshy fruit biology due to the
availability of an array of genetic and genomic resources. The basic knowledge gained from
tomato can often be applied to additional fruit crop species. Therefore, identification of genes
and their functional characterization in tomato will generate knowledge that can be utilized to
improve the production, yield, quality and nutritional value of additional fruit crops.
The cuticle is a hydrophobic protective layer present on all the aerial surfaces of
terrestrial plants. In fruit crops the cuticle provides structural integrity to the fruit along with
protection from uncontrolled water loss or uptake, mechanical damage, pathogens, herbivores
and thereby influences growth, ripening and post harvest quality, particularly in relation to fruit
cracking (Chu and Thompson, 1972; Emmons and Scott, 1997; Emmons and Scott, 1998; Matas
185

et al., 2004; Saladie et al., 2007; Dominguez et al., 2011). The presence of much thicker cuticle
in fruits compared to leaves of tomato (Nadakuduti et al., 2012) suggests its mechanistic role in
maintaining fruit integrity and physiological role in regulating post harvest water loss from the
astomatous fruit surface. Identifying the regulatory factors and transcriptional mechanisms
regulating fruit cuticle formation will lead to improved understanding of the role of cuticle in
fruit development and post harvest maintenance. It also provides opportunity for selecting
desirable cuticle traits for use in breeding programs targeting improvement of fruit quality.
Fruit color is a horticultural trait with large impact on the nutritional quality of the fruit.
Chloroplast development and chlorophyll accumulation in developing fruits are critical for fruit
photosynthesis, which contributes to up to 20% of fruit carbohydrates (Hetherington et al., 1998)
and is therefore important for determining the sugar, nutritional composition and flavor of the
ripe fruit (Powell et al., 2012). Thereby enhancing the photosynthetic competency of the fruit by
increasing the efficiency of chloroplast function or by increasing chloroplast number, represent
potential strategies for enhancing the nutritional value and flavor of fleshy fruits. The discovery
that KNOX genes regulate fruit-specific chloroplast development not only advances our scientific
knowledge of a basic process unique to plants but also creates the potential to identify novel
targets for selection in fruit crops that could be used for improving nutritional quality of fleshy
fruits.
In the US, consumption of processed tomatoes outweighs that of the fresh produce
(USDA-ARS 2006) and tomatoes are the highest consumed canned vegetable (National
Agricultural Statistics Service, 2005). Before processing and canning the fruit peel is generally
removed by various processes including steam, hot water, lye or cryogenic methods (Garcia and
Barrett, 2006). Removal of the fruit peel represents an essential component of tomato processing

186

that is important to the final quality of the product. However, this requires extra inputs in the
form of energy or chemicals such as lye that creates issues with waste disposal. As such, there
might be an advantage of the easy peeling phenotype of the ep mutant in the processing and
canning industry as fruit can be peeled without any pre-treatment. Once identified it may be
possible to introduce the ep mutant into commercial canning varieties to exploit the potential of
easy peeling during processing. However, the agricultural performance of such a trait would first
need to be explored as there is the potential negative impact of increased disease incidence in ep
fruit. As the ep mutant likely carries a deletion allele (Verkerk and Contant, 1969) it likely
represents a severe mutant allele but once the gene is identified, it may become possible to
isolate additional mutant alleles from EMS mutagenized populations that are less severe than the
original ep mutant allele. Alternatively, it may be possible to use the existing allele as a
heterozygote in F1 hybrids if an intermediate phenotype is observed. These approaches may
allow the fruits to be peeled easily without the associated disease penalty.
The long-term vision of the work described in this dissertation is to develop an improved
understanding of the fundamental biological processes that determine fruit development and
quality while identifying genes that may be useful for integration into tomato or additional fleshy
fruit crop breeding programs.

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