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CELL WALL MODIFICATIONS REGULATE FLOWER
DEVELOPMENT IN DENDROBIUM CRUMENATUM
YAP YOU MIN
[B. Sc. (Hons.), NUS]
A THESIS SUBMITTED
FOR THE DEGREE OF MASTER OF SCIENCE
DEPARTMENT OF BIOLOGICAL SCIENCES
NATIONAL UNIVERSITY OF SINGAPORE
2008
Acknowledgements
I would like to express my sincere gratitude to my supervisors, Dr. Ong Bee Lian and
A/P Loh Chiang Shiong, for their constant guidance and advice throughout the course of this
research.
I would also like to especially thank A/P Yeoh Hock Hin for providing the various
enzymatic substrates, Dr. Carol Han and Mr. Heng Mok Wei Dennis for their invaluable help
and advice regarding molecular work, Mr. Chong Ping Lee for his expertise in histology
work, Mr. Yap Wee Peng, Mr. Hee Kim Hor Daryl, Mr. Koh Teng Seah, Miss Ng Seow
Leng and Miss Lim Huiqin for their assistance in many ways.
Big thanks also goes to Daphne, June, Cipto, David and Sinteck for their help in one
way or another, and for providing much cheer and joy in the laboratory.
Lastly, I would like to thank my family and Chuanling for their constant love and
support.
ii
Contents
Page no.
Acknowledgements
Abstract
List of Tables
List of Figures
List of Abbreviations
ii
vi
viii
ix
xi
Chapter 1. Introduction
1
Chapter 2. Literature Review
2.1 Plant cell wall
2.1.1 Primary cell wall components
2.1.2 Bonds between cell wall components
4
4
4
6
2.2 Cell wall metabolism
2.2.1 Pectin demethylesterification
2.2.2 Pectin solubilisation
2.2.3 Cellulose metabolism
2.2.4 Hemicellulose metabolism
8
8
9
10
11
2.3 Cell wall hydrolases
2.3.1 Cellulase
2.3.2 Polygalacturonase
2.3.3 Pectin methylesterase
2.3.4 Exo-glycosidases
12
12
13
15
16
2.4 Flowers and their influences
18
2.5 Flowering physiology
2.5.1 Cell wall changes during flowering
2.5.2 Floral bud opening, flower longevity and their regulators
2.5.3 Flowering and the senescence programme
19
20
22
23
2.6 Orchids
2.6.1 Dendrobium crumenatum
24
25
Chapter 3. Materials and Methods
3.1 Plant material
27
27
3.2 Physical parameters analyses
27
iii
3.3 Microscopy
30
3.4 Electrolyte leakage
31
3.5 Cell wall compositional analyses
3.5.1 Preparation of EIR
3.5.2 Cellulose content
3.5.3 Total pectin content
3.5.4 Soluble pectin and hemicellulose contents
31
31
32
33
33
3.6 Cell wall enzyme analyses
3.6.1 Enzyme extraction
3.6.2 Cellulase assay
3.6.3 Polygalacturonase assay
3.6.4 Pectin methylesterase assay
3.6.5 Glycosidases assay
3.6.6 Soluble protein content
35
35
37
39
39
40
40
3.7 Gene expression profiling
3.7.1 Total RNA isolation
3.7.2 Estimation of RNA quality and quantity
3.7.3 Reverse transcription
3.7.4 PCR amplification
41
41
42
42
43
3.8 Controlling time of flower opening in D. crumenatum
3.8.1 General setup
3.8.2 Treatment
43
43
44
3.9 Statistical analysis
44
Chapter 4. Results
4.1 Growth changes
45
45
4.2 Changes in anatomy during development
45
4.3 Cell wall composition
4.3.1 EIR
4.3.2 Cellulose
4.3.3 Hemicelluloses
4.3.4 Total pectins
4.3.5 Soluble pectins
48
48
52
52
52
53
4.4 Activities of cell wall-based enzymes
4.4.1 Soluble proteins
4.4.2 Cellulase
56
56
58
iv
4.4.3 Polygalacturonase
4.4.4 Pectin methylesterase
4.4.5 β-galactosidase
4.4.6 β-glucosidase
4.4.7 β-mannosidase
4.4.8 β-xylosidase
61
61
63
65
65
68
4.5 Expression of cell wall-based enzyme gene transcripts
4.5.1 Total RNA integrity and quality
4.5.2 Optimization of PCR
4.5.3 Expression of β-GAL and PME during floral bud and flower development
68
70
70
74
4.6 Membrane stability
74
4.7 Control of flower opening
78
Chapter 5. Discussion
5.1 Cell wall changes related to D. crumenatum floral bud/ flower development
83
83
5.2 Model for cell changes accompanying D. crumenatum floral bud/ flower
development
89
5.3 Species-specific variations in cell wall modifications associated with flowering 91
5.4 Relationship between flowering and senescence
95
5.5 Effects of growth regulators on the control of flower opening
96
5.6 Further works
100
Chapter 6. Conclusion
101
Chapter 7. References
102
Appendix A:
117
Publication in Scientia Horticulturae
v
Abstract
The involvement of cell wall modifications, in particular, changes in cell wall components
and activities of cell wall-based enzymes, in regulating flower development in a sympodial
orchid, Dendrobium crumenatum, were investigated. Plants were subjected to cold treatments
to release floral buds from dormancy, and various parameters were investigated from young
floral bud stage till flower senescence. Anatomical studies demonstrated structural
disorganization in sepals and petals in developing floral buds. The packing and arrangement
of the cells were observed to become increasingly disorganized during flower opening and
flower senescence. Subsequent analysis of cell wall composition showed that the cell walls of
sepals and petals were modified extensively during floral bud development, flower opening
and flower senescence, as observed by the changes in the amounts of cellulose,
hemicelluloses and total pectins. Pectin solubilisation was also observed to commence during
early floral bud development. Of the tested cell wall-based enzymes, β-glucosidase
demonstrated the highest specific activity, followed by pectin methylesterase and βgalactosidase. Significant changes in the activities of the enzymes were also observed during
floral bud and flower development. The results indicated that cell wall modifications began
early in young floral buds, and regulated flower development. A model for cell wall
modifications, which involved loosening of the cellulose/hemicellulose and pectin networks,
in D. crumenatum was proposed. Furthermore, comparisons of the cell wall modifications in
D. crumenatum floral buds/ flowers to those in other species suggested the presence of
species-specific changes.
Throughout the development of D. crumenatum floral buds up till flower opening,
senescence hallmarks, such as the decrease in membrane stability, were observed. Attempts
vi
were made to generate floral buds that exhibited abnormal patterns of flowering for future
studies. This would allow comparisons of cell wall modifications (and other physiological
factors) between the normal and abnormal floral buds. Exogenous application of the plant
growth regulator, benzyladenine, was found to suppress flower opening and caused floral
buds to abort.
vii
List of Tables
Page no.
Table 1. Stages of floral bud development in D. crumenatum.
28
Table 2. Extraction procedures of cellulase and PG from floral buds of D.
crumenatum.
36
Table 3. Total and soluble pectins in EIR derived from D. crumenatum sepals at
various developmental stages.
54
Table 4. Total and soluble pectins in EIR derived from D. crumenatum petals at
various developmental stages.
55
Table 5. Quality of RNA obtained from D. crumenatum samples at various floral
bud/ flower developmental stages.
72
Table 6. Percentages of D. crumenatum floral buds that displayed full flower
opening one day after treatment.
81
Table 7. Percentages of D. crumenatum floral buds that displayed dormancy after
one day treatments, and percentages of dormany floral buds that
subsequently aborted two days after treatment.
82
Table 8. Summary of modifications of cell wall polysaccharides and cell wall
enzyme activities during floral bud development, flower opening and
flower senescence of D. crumenatum, carnation, sandersonia and
daylily.
94
viii
List of Figures
Page no.
Fig. 1
Dendrobium crumenatum (pigeon orchid).
26
Fig. 2
Separation of floral parts in D. crumenatum floral buds and flowers.
29
Fig. 3
Sequential extraction of pectins and hemicelluloses from EIR of D.
crumenatum floral buds.
34
Fig. 4
Characteristics of weight changes of sepals and petals of D.
crumenatum.
46
Fig. 5
Changes in anatomical features of D. crumenatum during flower
development.
47
Fig. 6
Anatomical changes during D. crumenatum floral bud development.
49
Fig. 7
D. crumenatum sepal and petal cell wall components, expressed on a
per floral part basis, at each developmental stage.
50
Fig. 8
D. crumenatum sepal and petal cell wall components, expressed on a
per gram fresh weight basis, at each developmental stage.
51
Fig. 9
D. crumenatum sepal and petal soluble protein contents at each
developmental stage.
57
Fig. 10
Optimisation of cellulase gel diffusion assay using commercial
cellulase.
59
Fig. 11
Gel diffusion assay for cellulase.
60
Fig. 12
Changes in pectin methylesterase activity in sepals and petals during
development of D. crumenatum floral buds.
62
Fig. 13
Changes in β-galactosidase activity in sepals and petals during
development of D. crumenatum floral buds.
64
ix
Fig. 14
Changes in β-glucosidase activity in sepals and petals during
development of D. crumenatum floral buds.
66
Fig. 15
Changes in β-mannosidase activity in sepals and petals during
development of D. crumenatum floral buds.
67
Fig. 16
Changes in β-xylosidase activity in sepals and petals during
development of D. crumenatum floral buds.
69
Fig. 17
Electrophoresis of total RNA from sepals and petals of D.
crumenatum.
71
Fig. 18
Gradient PCR amplification of β-TUB, β-GAL and PME transcripts.
73
Fig. 19
PCR amplifications of β-TUB, β-GAL and PME transcripts over 24,
26, 28, 30 and 32 cycles.
75
Fig. 20
Expression of β-GAL and PME transcripts at various developmental
stages of D. crumenatum floral buds/flowers.
76
Fig. 21
Membrane stability of sepals and petals of D. crumenatum at various
floral bud/flower developmental stages.
77
Fig. 22
General physical features of day 9 D. crumenatum floral buds after
treatments.
80
Fig. 23
Proposed model for cell wall modifications accompanying D.
crumenatum floral bud and flower development.
90
Fig. 24
Comparison of flower development events in carnation, sandersonia,
daylily and D. crumenatum.
92
x
List of Abbreviations
AOA
Ara
BA
CDTA
CMC
CSP
DNS
EDTA
EIR
GA
Gal
GalA
Gal-Fuc
GOD
MSI
PAR
PCD
PG
PGR
PME
Rha
SSP
TBA
Xyl
β-gal
β-glu
β-man
β-xyl
Aminooxyacetic acid
α-L-arabinose
Benzyladenine
trans-1,2-cyclohexanediamine-N,N,N’,N’-tetraacetic acid
Carboxymethylcellulose
CDTA-soluble pectins
Dinitrosalicylic acid
Ethylenediaminetetraacetic acid
Ethanol-insoluble residue
Gibberellic acid
α-D-galactose
D-galacturonic acid
β-D-galactosyl-α-L-fucose
Glucose oxidase/о-dianisidine
Membrane stability index
Photosynthetically active radiation
Programmed cell death
Polygalacturonase
Plant growth regulator
Pectin methylesterase
1,2-α-L-rhamnose
Na2CO3-soluble pectins
Tertiary-butanol
α-D-xylose
β-galactosidase
β-glucosidase
β-mannosidase
β-xylosidase
xi
Chapter 1. Introduction
Flowering is the first step of sexual reproduction in plants, and is a highly controlled
biological event in the life cycle of the angiosperms (Bernier et al. 1993; van Doorn and van
Meeteren 2003). Flowering and the eventual senescence of the flowers are also events of
commercial value, as they contribute to the visual quality and postharvest vase-life of the
flowers (O’Donoghue et al. 2002; Nell 2007). The cut flower trade has become a globalized
market, involving US$4.5 billion in international trade yearly, and with Singapore as one of
the major players in orchid export (Hew and Yong 2004; O’Donoghue 2006). Besides, with
the world’s increasing interest in ‘green buildings’ to aid energy efficiency and the
accompanying issue of using flowers for aesthetic benefits (Spala et al. 2008), it is important
to understand the biochemistry, physiology and genetics for flowering, longevity and
senescence (Nell 2007). However, publications on the study of tropical flowers are limited
and the few detailed studies focus mainly on flowers of temperate species.
There has been some focus on the possible involvement of cell wall modifications
and/or cell wall remodelling in the regulation of flowering (O’Donoghue 2006). Flowers that
have been studied include alstroemeria (Alstroemeria peruviensis), carnation (Dianthus
caryophyllus L.), daylily (Hemerocallis spp.) and sandersonia (Sandersonia aurantiaca
Hook.) (de Vetten and Huber 1990; Panavas et al. 1998; O’Donoghue et al. 2002; Wagstaff
et al. 2003). These studies also addressed the question on whether flowering could be a
process regulated by the senescence programme of the plant. Some supporting evidence
include the increase in oxidation of membrane components prior to flower opening in daylily
(Panavas and Rubinstein 1998), and the increasing trend of DNA laddering throughout petal
development in alstroemeria (Wagstaff et al. 2003). Studies on carnation and sandersonia
1
flowers demonstrated that the transitions of floral stages from opening floral bud to fully
mature flower till senescence were accompanied by changes in the levels of various cell wall
polymers, such as cellulose and pectins, and activities of cell wall-based enzymes (de Vetten
and Huber 1990; O’Donoghue et al. 2002). These observations were similar to the loss of
cell wall integrity in ripening fruits of carambola (Averrhoa carambola) and grapes (Vitis
vinifera) (Chin et al. 1999; Deng et al. 2005). In daylily flowers, analyses of cell wall
composition were not published, but reported changes in activities of cell wall-based
enzymes during flower development suggested the involvement of cell wall metabolism in
flowering (Panavas et al. 1998). While cellulase activity was detected in daylily flowers, it
was reported to be absent in sandersonia flowers, indicating the possibility of a speciesspecific variation in cell wall metabolism that regulates flowering (Panavas et al. 1998;
O’Donoghue et al. 2002).
In the above-mentioned studies on the possible involvement of cell wall
modifications and cell wall remodelling in regulating flowering, cell wall changes were
compared only between stages of mature bud (just prior to opening), opening flower, mature
flower, wilting flower and senesced flower (de Vetten and Huber 1990; O’Donoghue et al.
2002; Wagstaff et al. 2003). To fully understand if and/ or how flowering is regulated by a
senescence programme that has already started, investigating the physiological, biochemical
and molecular changes occurring throughout the development of a newly-induced young
floral bud till flower senescence would be advantageous.
Few studies on the physiology of flowering in tropical orchids have been conducted
to date. As orchid cultivation continues to be a highly profitable commercial market (Hew
and Yong 2004), characterization of tropical orchid flowering is of paramount importance.
2
Dendrobium crumenatum (Swartz), also known as the pigeon orchid, is a common native
epiphytic orchid species of South-east Asia. The floral buds of D. crumenatum exhibit
dormancy, and can be induced to resume growth and development by cold-induction,
culminating into the opening of the flowers exactly nine days after (Holttum 1953; Corner
1988). Flower opening in D. crumenatum is a rapid and short process, taking about 4 hours
from the onset of floral bud crack, achieving full flower opening before dawn (Yap 2006).
The flowers are short-lived, lasting for only 24 h under natural conditions, before senescence
sets in (Tan and Hew 1993). The synchronized and predictable pattern of floral bud
development in D. crumenatum, together with its short life cycle, makes the pigeon orchid an
ideal system to study the control of flowering.
In this study, anatomical changes in sepals and petals were studied over various
stages throughout the development of newly-induced floral buds till flower senescence in D.
crumenatum. The levels of cell wall components such as cellulose, hemicellulose and
pectins, activities of various cell wall-based enzymes, and their corresponding expression
levels of the gene transcripts were also followed throughout development. We aim to use all
these data to help us understand the involvement of cell wall modifications and remodelling
in the regulation of flowering in tropical orchids.
We also aimed to develop a system to obtain floral buds that exhibit abnormal flower
opening patterns. Attempts to delay flower opening, using plant growth regulators, were
made in mature D. crumenatum floral buds. Such system would be useful for future studies
that may involve analysing and comparing cell wall modifications between floral buds
showing normal and abnormal opening patterns.
3
Chapter 2. Literature Review
2.1 Plant cell wall
Each plant cell comprises a specialised and complex cell wall that serves many functions
(Cosgrove 1999). The cell walls provide structural support and maintain the shape of the
cells, act as a protective barrier against water loss, pathogens, and other mechanical and
environmental stresses, take part in cell-cell communication and interaction by carrying
surface signalling molecules, and act as a storage organ for carbohydrates, proteins and
various other materials (Cosgrove 1999; Carpita and McCann 2000). The high mechanical
strength and rigidity of the cell walls allowed plants to become some of the largest organisms
on Earth. Yet, the cell walls remain extensible, allowing plant cells to grow until the cells
cease growth (Cosgrove 1999). The dynamic structure of the cell wall is important in
regulating cell expansion and cell growth. For example, cell wall loosening is a pre-requisite
for the incorporation of newly synthesized wall polymers during cell expansion and cell
growth (Carpita and McCann 2000). Plant cell walls also have important roles in controlling
fruit ripening. The degree of cell wall disassembly and/or cell wall weakening in fruits has
been shown to regulate the time of ripening and the texture of a variety of fruits (Brummell
2006). Cell wall modifications in petals have also been linked to flowering (O’Donoghue
2006).
2.1.1 Primary cell wall components
The primary cell wall is a complicated matrix, composed of various polymers made up of
polysaccharides, proteins and some phenolics (Carpita and McCann 2000; Brummell and
Harpster 2001; Brummell 2006; Liepman et al. 2007). A highly hydrated complex, the
4
primary cell wall also contains various aromatic substances, dissolved solutes and ions, and
soluble proteins including enzymes (Brummell 2006).
Cellulose is the most abundant plant polysaccharide and acts as the principal scaffold
in plant cell walls (Carpita and McCann 2000). Cellulose microfibrils are composed of 1,4-βD-glucan
chains assembled together by extensive hydrogen bonding, resulting in long, rigid,
inextensible fibres. The cellulose microfibrils have a crystalline internal region that excludes
water, and an amorphous outer layer that interacts with other matrix molecules (Pauly et al.
1999; Carpita and McCann 2000; Brummell 2006; Liepman et al. 2007).
Another main component of the cell wall is hemicellulose (also known as crosslinking glycan), which can hydrogen bond to cellulose microfibrils, thus forming a network
between various microfibrils (Carpita and McCann 2000). Predominantly made up of neutral
sugars, hemicelluloses are neutral or weakly acidic. There are three major types of
hemicellulose. The first, xyloglucan, is the most abundant. Similar to cellulose, xyloglucan
comprises of a 1,4-β-D-glucan backbone, but has regularly spaced α-D-xylose (Xyl) side
chains (on three consecutive glucose residues out of four). The xylose side chains may also
be extended with β-D-galactosyl-α-L-fucose (Gal-Fuc) or α-L-arabinose (Ara). Another major
hemicellulose is xylan, which has a backbone comprising of 1,4-β-linked xylopyranosyl
units. Another form of xylan is arabinoxylan, which has a backbone consisting of 1,4-β-Dxylan, with occasional α-L-arabinose substitutions. The third major hemicellulose is
(galacto)glucomannan, comprising of alternating regions of 1,4-β-D-glucan and 1,4-β-Dmannan in approximately equal amounts. Single units of terminal α-D-galactose (Gal) are
also occasionally found on galactoglucomannan (Carpita and McCann 2000; Brummell and
Harpster 2001; Brummell 2006; Minic and Jouanin 2006; Liepman et al. 2007).
5
The cell wall is also a pectin rich construct. Pectins belong to a class of
polysaccharides that can be linear or branched, highly hydrated and rich in D-galacturonic
acid (GalA) residues. Some functions of pectins include regulating wall porosity (Baron-Epel
et al. 1988) and cell-cell adhesion at the middle lamella (Pena and Carpita 2004). Due to their
ability to control wall porosity, pectins may also affect cell wall modifications by regulating
the access of cell wall enzymes to their respective substrates in the matrix (Carpita and
McCann 2000; Brummell 2006). One of the fundamental constituents of pectins is
homogalacturonan, which has a backbone of 1,4-α-D-GalA. Pectins may exist as linear,
unbranched
homopolymers
of
1,4-α-D-GalA,
or
exist
as
structurally
modified
homogalacturonans, such as xylogalacturonan, which has a homogalacturonan backbone with
single Xyl side chains. Another modified homogalacturonan is rhamnogalacturonan II, which
has a homogalacturonan backbone with highly conserved side chains, consisting of a
diversity of neutral sugars. Pectins may also exist as rhamnogalacturonan I.
Rhamnogalacturonan I has a backbone of alternating 1,2-α-L-rhamnose (Rha) and GalA
disaccharide units, and may also possess linear or branched arabinan and galactan side chains
(Carpita and Gibeaut 1993; Carpita and McCann 2000; Brummell 2006; Liepman et al.
2007). In brief, the primary cell wall consists of homogalacturonan, rhamnogalacturonan I,
rhamnogalacturonan II, hemicellulose and cellulose, with an almost equal distribution of
each component, whereas the middle lamella consists of mainly homogalacturonan and
structural proteins (Brummell 2006).
2.1.2 Bonds between cell wall components
The various cell wall components are linked by a variety of bonds (Brummell 2006).
Hydrogen bonds occur extensively between cellulose microfibrils and also help to bind
6
xyloglucan to cellulose. Homogalacturonan molecules are attached to each other by ionic
calcium bridges, and pectin molecules are bound to each other, or to other pectins,
hemicelluloses or phenolic molecules via ester linkages (Fry 1986; Carpita and Gibeaut
1993; Iiyama et al. 1994; Rose and Bennett 1999; Carpita and McCann 2000; Liepman et al.
2007). Covalent linkages have also been reported between homogalacturonan and
rhamnogalacturonan II (Vincken et al. 2003), between xyloglucan and arabinan or galactan
side chains of rhamnogalacturonan I (Thompson and Fry 2000; Popper and Fry 2005),
between rhamnogalacturonan I and extensin (Qi et al. 1995), and between various structural
proteins and phenolics (Fry 1986). Besides being chemically bonded to one another, the
various cell wall components may also be attached to each other by physical means, via
entanglement, for instance (Brummell 2006). For example, rhamnogalacturonan I side chains
are wound around cellulose microfibrils, creating a pectin network that is interlocked with
the cellulose-hemicellulose network (Vincken et al. 2003; Zykwinska et al. 2005).
Due to the various bonds present in the cell wall matrix, the study of cell wall
composition requires specific chemical treatments to release the respective cell wall
components (Brummell 2006). The chemical treatments include chelating agents such as
trans-1,2-cyclohexanediamine-N,N,N’,N’-tetraacetic
acid
(CDTA)
or
ethylenediaminetetraacetic acid (EDTA) to extract ionically bound pectins, sodium carbonate
to extract pectins held by ester bonds, weak alkali such as 1 M KOH to extract loosely
attached hemicelluloses, and strong alkali such as 4 M KOH to extract hemicelluloses held
tightly by hydrogen bonds (Brummell 2006).
7
2.2 Cell wall metabolism
Various regulated cell wall architecture changes occur with the development of plants
(Carpita and McCann 2000). Fruit ripening is one developmental event whereby many
changes occur in the cell walls, resulting in the final texture of the fruit (Brummell 2006).
Consequently, the majority of the information on cell wall metabolism is derived mainly
from
fruits.
Some
of
the
cell
wall
modifications
involved
include
pectin
demethylesterification, pectin solubilisation, and metabolism of cellulose and hemicellulose
(Carpita and McCann 2000; Brummell and Harpster 2001; Brummell 2006).
2.2.1 Pectin demethylesterification
During pectin synthesis, pectins are polymerised in the cis Golgi, and are subsequently
methylesterified in the medial Golgi (Goldberg et al. 1996). The methylesterified pectins
may also be substituted with side chains in the medial Golgi cisternae (Goldberg et al. 1996).
Thus, pectins are secreted into plant cell walls in highly methylesterified forms (Carpita and
McCann 2000; Micheli 2001). The degree of methylesterification, however, decreases with
development and has been shown to be a crucial physiological change during fruit ripening
(Brummell et al. 2004), microsporogenesis and pollen tube growth (Wakeley et al. 1998;
Futamura et al. 2000), seed germination (Ren and Kermode 2000), and hypocotyl elongation
(Bordenave and Goldberg 1993). Roy et al. (1992) demonstrated that the time of onset and
area of the cell wall to undergo pectin demethylesterification were tightly regulated in tomato
(Lycopersicon esculentum). Highly methylesterified pectins became increasingly less
methylesterified as ripening of tomatoes progressed, and the demethylesterification process
started in the middle lamella, spreading throughout the rest of the cell wall (Roy et al. 1992).
8
The removal of methylester groups from pectin results in negatively charged
carboxylic groups (Grignon and Sentenac 1991). These charged surfaces may be involved in
regulating pH and ion balance, in turn, affecting the activity of cell wall hydrolases (Chun
and Huber 1998; Almeida and Huber 1999). The charged surfaces resulting from
demethylesterification of pectins may also affect the movements of charged molecules, such
as proteins, within the cell wall matrix (Grignon and Sentenac 1991). In the presence of
calcium, the demethylesterified charged pectate molecules can aggregate and bind to one
another via calcium cross-links, forming calcium-pectate gels which can increase stiffness of
the cell wall (Jarvis 1984).
2.2.2 Pectin solubilisation
One of the major changes in cell wall pectins with the development of plants is the increasing
solubilisation of pectins, as commonly observed during the ripening of fruits (Brummell and
Harpster 2001). Pectin solubilisation is usually measured as the increase in ease of
extractability of pectins by various extractants, and can be extrapolated to bond changes
within the cell wall matrix (Brummell and Harpster 2001). In watermelon (Citrullus lanatus),
the amounts of water-soluble and chelator-soluble pectins increased at the expense of sodium
carbonate-soluble pectins during ripening (Rose et al. 1998). In tomato and avocado (Persea
americana), increases in amounts of water-soluble pectins were observed in conjunction with
decreases in the amounts of sodium carbonate-soluble pectins, but changes in amounts of
chelator-soluble pectins were absent (Carrington et al. 1993; Wakabayashi et al. 2000). The
increases in water-soluble and/or chelator-soluble pectins were attributed to the increasing
proportions of pectins that were more weakly attached to the cell wall matrix.
9
Pectin demethylesterification has been proposed as a possible cause for pectin
solubilisation (Brummell 2006). The resultant regions of negatively-charged groups during
demethylesterification could cause electrostatic repulsion between negatively-charged
molecules, detaching the pectins that were weakly bound to the cell wall (Grignon and
Sentenac 1991). The loss of arabinan and galactan side chains from rhamnogalacturonan I
has also been suggested to cause pectin solubilisation (Brummell 2006). The arabinan and
galactan side chains firmly bind pectins to the cell wall via covalent linkages, hydrogen
bonds or physical entanglement (Popper and Fry 2005; Zykwinska et al. 2005). Thus the loss
of the side chains could cause loosening of the pectins. In papaya (Carica papaya), the
presence of the galactan degrading enzyme, β- galactosidase, also caused increased pectin
solubilisation (Ali et al. 1998). Pectin solubilisation is believed to result in cell wall swelling
(Redgwell et al. 1997). This would indirectly cause changes in the movement of cell wall
enzymes through the wall matrix, increasing their accessibility to their respective substrates
(Brummell 2006).
2.2.3 Cellulose metabolism
Due to the insolubility of cellulose in standard solvents, and its highly susceptible nature to
hydrolysis in harsh solvents, the quantification of cellulose changes during development of
plants is a challenging procedure. Consequently, little has been published regarding cellulose
degradation. As the main component of the cell wall matrix, cellulose is expected to undergo
distinct alterations during cell wall changes (Fischer and Bennett 1991; Rose and Bennett
1999). Cell wall cellulose content decreased during ripening of grapes (Deng et al. 2005).
However, during the ripening of pear (Pyrus communis), tomato and avocado, cellulose
levels remained constant or even increased slightly (Gross and Wallner 1979; Ahmed and
10
Labavitch 1980; Maclachlan and Brady 1994; Sakurai and Nevins 1997). The resistance of
cellulose towards enzymatic degradation in most fruits reflects the classical description of
cellulose microfibrils as structurally stable and highly crystalline (Rose and Bennett 1999). It
has also been suggested that cellulose metabolism is not a major feature of cell wall
modifications during plant development, specifically, during fruit ripening (Brummell 2006).
2.2.4 Hemicellulose metabolism
It has been widely accepted that cellulose microfibrils are coated with hemicelluloses,
particularly xyloglucan, on their surfaces and are further cross-linked by xyloglucan chains,
thus creating a three-dimensional cellulose-xyloglucan network (Fischer and Bennett 1991;
Rose and Bennett 1999; Carpita and McCann 2000). Hemicellulose metabolism would, thus,
significantly affect the extent of the cellulose-xyloglucan network and the primary cell wall
structure (Fischer and Bennett 1999; Carpita and McCann 2000; Brummell 2006).
Hemicellulose content has been shown to decrease during the ripening process in a variety of
fruits such as tomato, strawberry (Fragaria ananassa), muskmelon (Cucumis melo),
capsicum (Capsicum annum), pepino (Solanum muricatum), carambola, grapes, boysenberry
(Rubus idaeus x Rubus ursinus) (Huber 1983a, 1984; McCollum et al. 1989; Sethu et al.
1996; O’Donoghue et al. 1997; Chin et al. 1999; Deng et al. 2005; Vicente et al. 2007).
The breakdown of hemicelluloses has been suggested to be a major contributor to
reduced cell wall turgidity, resulting in the softening of cell walls and possible cell wall
expansion (Fischer and Bennett 1999; Brummell 2006). Relaxation of the cellulosexyloglucan network brought about by xyloglucan breakdown could also cause cell wall
swelling (Brummell 2006), which affects pectin solubilisation as mentioned previously.
11
2.3 Cell wall hydrolases
The various cell wall modifications are the results of the actions of a range of cell wallmodifying enzymes, the activities of which vary with development. The activities of cell wall
hydrolases and the expression of their corresponding genes have thus been intensely studied
to fully understand cell wall modifications (Fischer and Bennett 1991; Brummell and
Harpster 2001; Minic and Jouanin 2006). To cope with the complex cell wall polymers, the
activities of the cell wall localised or plasma membrane bound enzymes are very diverse
(Minic and Jouanin 2006). The majority of the information on cell wall hydrolases in plants
is derived from studies on Arabidopsis and various fruits.
Various families of cell wall hydrolases, such as glycosidases (also known as
glycoside hydrolases) and carbohydrate esterases, have been shown to participate in cell wall
modifications (Minic and Jouanin 2006). Glycosidases are enzymes that catalyse the
hydrolysis of the glycosidic linkages between two or more carbohydrates or between a sugar
moiety and a non-sugar moiety (Davies and Henrissat 1995). Cellulase, polygalacturonase, βglucosidase, β-galactosidase, β-mannosidase and β-xylosidase belong to the glycosidase
family (Minic and Jouanin 2006; Minic 2008). Carbohydrate esterases are enzymes that
catalyse the removal of non-carbohydrate groups on substituted polysaccharides. For
example, pectin methylesterase belongs to this family of enzymes, and catalyses the removal
of methyl groups from polysaccharides (Micheli 2001; Minic and Jouanin 2006).
2.3.1 Cellulase
Modifications in cell wall glucans occur due to the actions of endo-1,4-β-glucanase (EC
3.2.1.4), or more commonly referred to as cellulase (Wood and Bhat 1988; Brummell and
Harpster 2001). Cellulase catalyses the hydrolysis of linkages of 1,4-β-D-glucan chains
12
adjacent to unsubstituted residues (Brummell and Harpster 2001). Substrates of plant
cellulase include carboxymethyl cellulose, amorphous cellulose and xyloglucan, and to a
lesser extent, crystalline cellulose (Ohmiya et al. 1995; Molhoj et al. 2001).
Cellulase activity has been detected in a variety of fruits, although the amount and
pattern of change vary considerably. During ripening of banana (Musa acuminata) and
pawpaw (Asimina triloba), cellulase activity (units mg protein-1) increased (Lohani et al.
2004; Koslanund et al. 2005); in carambola, guava (Psidium guajava), grapes and
boysenberry, cellulase activity (units gFW-1) also increased during ripening (Chin et al. 1999;
Abu-Bakr et al. 2003; Deng et al. 2005; Vicente et al. 2007). There was however no changes
in levels of cellulase activity (units mg protein-1) during ripening in capsicum (Sethu et al.
1996), and the level of cellulase activity (units gFW-1) decreased during ripening in apple
(Malus domestica) (Goulao et al. 2007). The action of cellulase on xyloglucan may cause the
breakdown of the cellulose-xyloglucan network, and has been suggested to be a mechanism
for fruit softening (Rose and Bennett 1999). However, molecular studies demonstrated that
suppressing expression of the mRNA of the cellulase genes, LeCel1 and LeCel2 in tomato,
and overexpression of CaCel1 in pepper did not substantially affect the ripening processes
(Brummell and Harpster 2001).
2.3.2 Polygalacturonase
Polygalacturonases (PGs) are cell wall-based enzymes that catalyse the hydrolytic cleavage
of galacturonide linkages in pectins (Fischer and Bennett 1991; Brummell and Harpster
2001). Both exo- and endo-acting types of PGs have been identified and characterized in
fruits (Hadfield and Bennett 1998). Exo-PG (EC 3.2.1.67) removes GalA residues from the
non-reducing ends of polygalacturonic acid, while endo-PG (EC 3.2.1.15) cleaves
13
polygalacturonic acid at random (Brummell and Harpster 2001). Endo-PG, rather than exoPG, has been correlated with cell wall modifications such as pectin degradation (Huber
1983b; Fischer and Bennett 1991; Hadfield and Bennett 1998). Consequently, the majority of
studies on cell wall changes focus on endo-PG, and the enzyme will be referred to hereafter
as PG.
The main substrates for PGs are the homogalacturonans that are secreted into plant
cell walls in highly methylesterified forms, which must be de-esterified before the enzyme
can hydrolyse them (Jarvis 1984; Carpita and Gibeaut 1993; Minic and Jouanin 2006). PG
has been suggested to be a key enzyme in cell wall modifications during fruit ripening and
softening (Fischer and Bennett 1991; Brummell and Harpster 2001; Minic and Jouanin
2006). During ripening of papaya, tomato, carambola, guava and grapes, high levels of PG
activity (units mg protein-1) were reported (Lazan et al. 1989; Chin et al. 1999; Abu-Bakr et
al. 2003; Deng et al. 2005); increase in PG activity (units gFW-1) was also observed in
bananas (Lohani et al. 2004). In other fruits such as strawberry and apple, PG activity were
reported to be absent, but PG activity and/or mRNA expression were subsequently detected
(Hadfield and Bennett 1998).
Using transgenic methods, PG was shown to be a major player in regulation of pectin
solubilisation in fruits. In ripening-impaired tomato fruits containing the rin mutation,
reduced accumulation of PG mRNA was observed, in conjunction with reduced PG activity
and pectin solubilisation (Della Penna et al. 1987, 1989; Seymour et al. 1987; Knapp et al.
1989). In the ‘rescue’ experiments, PG activity and the level of pectin solubilisation were
almost similar to those of the wild-type (Della Penna et al. 1990).
14
2.3.3 Pectin methylesterase
Pectin methylesterase (PME; EC 3.1.1.11) is an enzyme that contributes to the degradation of
pectins (Minic and Jouanin 2006). It acts by catalysing the removal of methyl groups from
the C6 position of GalA residues of high molecular weight pectins (Fischer and Bennett
1991; Brummell and Harpster 2001; Micheli 2001). The demethylesterification of pectins
releases acidic pectins and methanol as products, and causes changes in the pH and charge of
cell walls (Carpita and Gibeaut 1993; Stephenson and Hawes 1994; Micheli 2001). PME has
been suggested to be an important regulator for various cell wall modifications related to
pectin demethylesterification and pectin solubilisation, which had been previously described.
In carambola, guava, grapes and boysenberry fruits, activity of PME (units gFW-1) increased
throughout fruit development (Chin et al. 1999; Abu-Bakr et al. 2003; Deng et al. 2005;
Vicente et al. 2007); in capsicum, banana and pawpaw fruits, activity of PME (units mg
protein-1) demonstrated decreases upon full ripening (Sethu et al. 1996; Lohani et al. 2004;
Koslanund et al. 2005).
The mode of action of PME was previously described to be dependent on the pH of
the PMEs. Acidic PMEs resulted in random demethylesterification of pectins, and alkaline
PMEs resulted in linear (along the chain) demethylesterification of pectins (Markovic and
Kohn 1984). However, more recent studies demonstrated that the action pattern of PMEs is
much more complicated, and may be regulated by many factors such as pH and the degree of
methylesterification of the pectins (Catoire et al. 1998; Denes et al. 2000).
PME has been studied in great detail in tomato, and has been shown to consist of at
least four genes, some of which are highly homologous (Harriman et al. 1991; Hall et al.
1994; Turner et al. 1996; Gaffe et al. 1997). During the ripening of tomatoes, PME protein
and activity increased throughout and then declined slightly upon full ripening (Harriman et
15
al. 1991; Tieman et al. 1992). However, the accumulation of PME mRNA demonstrated an
opposite trend, decreasing as ripening progressed, and it was suggested that the nonsynchronised patterns were due to the quantification of a composite of PME proteins of two
or more highly homologous genes (Harriman et al. 1991; Brummell and Harpster 2001).
2.3.4 Exo-glycosidases
There is a variety of glycosidases because of the structural and functional diversity of the
polysaccharides and oligosaccharides (Davies and Henrissat 1995; Minic 2008). Some
examples of exo-glycosidases include β-galactosidase (β-gal; EC 3.2.1.23), β-glucosidase (βglu; EC 3.2.1.21), β-mannosidase (β–man; EC 3.2.1.25) and β-xylosidase (β-xyl; EC
3.2.1.37), and all have been detected during fruit development in mango (Mangifera indica),
capsicum, asparagus (Asparagus officinalis), carambola and boysenberry (Ali et al. 1995;
Sethu et al. 1996; O’Donoghue et al. 1998; Chin et al. 1999; Vicente et al. 2007, Minic
2008).
β-gal, which catalyses the removal of β-D-galactosyl residues from β-D-galactosides
has been relatively well-studied (Brummell and Harpster 2001). The loss of Gal in cell walls
is a common feature during fruit ripening, and it has been suggested that β-gal acts by
hydrolyzing the Gal residues from the sidechains of pectins, causing changes in the structure
of the pectin network (O’Donoghue et al. 1998; Brummell and Harpster 2001). β-gal exists
in at least three isoforms in tomato and mango (Pressey 1983; Ali et al. 1995). In tomato, βgal is encoded by at least seven genes TBG1 – TBG 7 (Smith and Gross 2000). Transcripts of
the seven genes exhibited differential expression during fruit development, and only TBG4
mRNA was significantly reduced in ripening-impaired rin and nor mutants (Smith and Gross
2000).
16
β-glu completes the breakdown of glucans by catalysing the hydrolysis of
oligosaccharides to release glucose (Wood and Bhat 1988; Hrmova and Fincher 2001). In
barley (Hordeum vulgare), β-glu demonstrates broad substrate specificity, hydrolysing βglucans, β-oligoglucosides and xyloglucan (Hrmova and Fincher 1998). Most studies on βglu focus on β-glucans as substrates, and little is known as to how β-glu participates in
remodelling of the cell wall structure (Hrmova and Fincher 2001). It has been suggested that
the ‘real’ substrate for β-glu is xyloglucan, thus affecting the cellulose-xyloglucan network
(Rose and Bennett 1999; Hrmova and Fincher 2001).
β-man is involved in the degradation of galactoglucomannans, by catalysing the
removal of Gal on the sidechains of manno-oligosaccharides (Minic and Jouanin 2006). The
activity of β-man in the cell walls of germinating seeds of monocotyledons, such as Phoenyx
dactylifera, has been investigated and it was suggested that the enzyme is involved in the
mobilization of galactoglucomannan in seeds (Buckeridge et al. 2000).
β-xyl takes part in the degradation of xylan and arabinoxylan (Minic and Jouanin
2006). The enzyme is identified as a key enzyme for the complete breakdown of xylan by
catalysing the hydrolysis of xylo-oligosaccharides from the non-reducing ends, and releasing
xylose (Minic and Jouanin 2006; Minic 2008). β-xyl has also been shown to be involved in
the degradation of arabinan side chains during hydrolysis of rhamnogalacturonan I (Minic et
al. 2004; Minic and Jouanin 2006).
Compared to endo-glycosidases (cellulase and PG) that break load bearing cross-links
in the cell wall, exo-glycosidases appear to cause much less significant effects as they
catalyse the removal of single glycosyl residues from polysaccharide chains (Rose and
Bennett 1999; Hrmova and Fincher 2001). However, it has been proposed that exo-
17
glycosidases may still participate in cell wall remodelling as side chain removal increases the
accessibility and availability of substrate sites for endo-acting enzymes (Rose and Bennett
1999). Also, actions of exo-glycosidases on side chains of xyloglucans may affect the
binding affinity of xyloglucan to cellulose microfibrils, hence disrupting the cellulosexyloglucan network (Rose and Bennett 1999).
2.4 Flowers and their influences
Flowering is a critical event in the life-cycle of angiosperms, allowing for the reproduction of
these plants. A complete flower consists of sepals and petals (together, they form the perianth
of the flower), gynoecium and androecium, which are the essential organs for sexual
reproduction in the higher plants (Bernier et al. 1993; Burger 2006; O’Donoghue 2006).
Biologically, sepals and petals play important roles by protecting immature reproductive
structures, then providing attraction and accessibility required for pollination to occur
(O’Donoghue 2006).
Flowers of various plants are also highly prized objects of beauty, and can be
considered as commercially valuable and luxurious commodities incorporated into everyday
lives (Hughes 2000; O’Donoghue 2006). The perishable nature of flowers, however, poses as
a constant setback for the horticulture industry (O’Donoghue 2006; Nell 2007). To help
prolong postharvest life of flowers, there are a variety of chemical preservative treatments,
including silver thiosulfate, 8-hydroxyquinoline citrate and/ or sucrose (Redman et al. 2002).
However, cold storage/ precooling/ refrigeration remains as the most common and
recognised form of postharvest treatment, as it effectively lowers the rate of plant metabolic
processes and the rate of microbial growth (Geertsen 1990; Sun and Brosnan 1999, 2001;
18
Redman et al. 2002; van Meeteren 2007). Disadvantages of precooling techniques include its
contribution to global warming, high energy demands, high costs, and the intolerance of
certain species of flowers towards cold (Redman et al. 2002; van Meeteren 2007; Kim and
Infante Ferreira 2008).
The importance of flowers has also been readdressed in recent years, in conjunction
with the world’s increasing interest in sustainability (Bartlett 1997; Espinosa et al. 2008;
Spala et al. 2008). For example, to help minimize energy consumption of buildings, ‘green
buildings’ or ‘green roofs’ have been developed. These structures make use of green plants to
provide shade for the buildings, control temperature and humidity, mitigate the greenhouse
effect, filter pollutants and mask noise, thus creating buildings that are energy efficient
(Spala et al. 2008). For aesthetic reasons, flowers are important components, and flower
physiology and flowering seasons have become essential criteria during plant selection
(Spala et al. 2008).
2.5 Flowering physiology
Flowering is considered a multifactorial process (Bernier 1988), comprising of two distinct
processes: floral initiation (floral induction), referring to the transition from vegetative to
reproductive development, and subsequent floral development, referring to the development
from floral bud to mature floral bud to mature flower and finally to senesced flower (Hew
and Yong 2004). In this thesis, the term ‘flowering’ will be used to define the processes
involved in floral development, from floral bud till senesced flower stages.
There are great variations among the angiosperms in the manner, timing, and
physiology of flowering (van Doorn and van Meeteren 2003; O’Donoghue 2006). In tulips
19
(Tulipa genesriana), flower opening is due to movements of the petal lamina as mesophyll
cells expand with temperature (Wood 1953). On the other hand, flower opening in Ipomoea
is due to movements of the midrib rather than the petal lamina (Kaihara and Takimoto 1981).
Flower opening in Portulaca occurs in the day (Ichimura and Suto 1998), while flower
opening in Oenothera lamarkiana occurs at night (Saito and Yamaki 1967). Physiologically,
it had been shown that carbohydrate metabolism (particularly the breakdown of storage
carbohydrates into soluble sugars) and water relations affecting cell wall turgor are involved
in regulating flowering in many species of flowers, such as alstromeria (Alstromeria
peregrine) (Collier 1997), rose (Rosa) (Evans and Reid 1988), daylily (Hemerocallis spp.)
(Bieleski 1993), lily (Lilium hybrid) (Bieleski et al. 2000), Campanula rapunculoides
(Vergauwen et al. 2000) and Capparis spinosa L. (Rhizopoulou et al. 2006).
Hormonal regulation is another influential mechanism of flowering. For example,
supplementing tulip flowers with gibberellin4+7 (GA4+7) plus benzyladenine (BA) helped to
promote longevity of the flowers (Kim and Miller 2008), exogenous ethylene caused delayed
flower opening in rose (Tan et al. 2006) and exogenous abscisic acid (ABA) resulted in
premature senescing of daylily flowers (Panavas et al. 1998). Changes in petal cell walls
have also been suggested to be involved in controlling flowering (van Doorn and van
Meeteren 2003; O’Donoghue 2006). Information on this topic is, however, very much
limited when compared to the other physiological processes mentioned above.
2.5.1 Cell wall changes during flowering
Only a few species of flowers have been studied on the influence of cell wall changes
in regulating flowering, and these include carnation (Dianthus caryophyllus L.), sandersonia
(Sandersonia aurantiaca Hook.) and daylily. Cell wall changes that occur in the flower
20
petals include alterations in cell wall compositions, activities of cell wall enzymes, and
expression of cell wall-related genes (O’Donoghue 2006). Modifications in petal cell walls
are necessary to provide the flexibility required during dramatic floral bud and flower growth
(van Doorn and van Meeteren 2003; O’Donoghue et al. 2005).
Comparison between the flowers shows certain similarities and differences in cell
wall changes associated with flowering. In carnation flowers, full flower opening was
accompanied by increases in contents of cell wall cellulose, total pectins, chelator-soluble
pectins, carbonate-soluble pectins and neutral sugars. Upon senescence of the flowers, all of
these cell wall components, except chelator-soluble pectins, decreased in content (de Vetten
and Huber 1990; de Vetten et al. 1991). While PG activity was absent in opened and
senesced flowers, activities of β-glu and β-gal were detected in senescing flowers (de Vetten
et al. 1991).
As with carnation flowers, sandersonia flowers also exhibited increase in contents of
cellulose, total pectins and neutral sugars during flower opening, and with the exception of
neutral sugars, the contents increased further during senescence (O’Donoghue et al. 2002).
Levels of chelator-soluble and carbonate-soluble pectins remained unchanged during flower
opening, and subsequently decreased upon senescence of the flowers. Similar to carnation
flowers, PG activity was also not detected in sandersonia flowers. Cellulase activity was also
reported to be absent in sandersonia flowers. The level of β-gal activity was unchanged
during flower opening in sandersonia flowers, and subsequently, increased during flower
senescence. In the same study, PME activity was found to increase during flower opening
and decreased upon flower senescence (O’Donoghue et al. 2002). Three genes that putatively
encode β-gal (SaGAL1, SaGAL2, and SaGAL3) have also been reported, and it was found
21
that all three genes were expressed during the onset of flower senescence (O’Donoghue et al.
2005).
Analysis of cell wall composition was not reported for daylily flowers, but analyses
on cell wall enzyme activities of daylily flowers indicated the participation of cell wall
changes during flower opening. Cellulase and PME activities increased during floral bud
development, and decreased upon senescence (Panavas et al. 1998). Unlike carnation and
sandersonia flowers, PG activity was detected in daylily flowers and was reported to increase
during senescence (Panavas et al. 1998).
In Arabidopsis flowers, although cell wall compositional changes were not reported,
cell wall enzyme activity assays and gene expression studies supported that cell wall
modifications were regular features of flower opening (O’Donoghue 2006). PME activities
were detected in mature Arabidopsis flowers, and at least one PME gene was shown to be
strongly expressed in the flowers (Micheli et al. 1998; Francis et al. 2006). Five PG genes
and 14 PME genes were reported to be differentially expressed in floral bud clusters (Imoto
et al. 2005).
2.5.2 Floral bud opening, flower longevity and their regulators
There are various endogenous regulators that control flower opening, and hormones are one
example (van Doorn and van Meeteren 2003). Some commonly investigated hormones or
plant growth regulators (PGRs) include the cytokinin, benzyladenine (BA), and gibberellins
(GAs) (Bernier 1988). Although various studies have been conducted to determine the effects
of BA and GA on flowering, generalizations of the effects of the PGRs cannot be applied
across for all plant species because some of the PGRs are present in supra- or sub-optimal
amounts (Cleland 1982).
22
Exogenous applications of BA have been shown to suppress flower opening, flower
wilting and senescence in roses, petunia and Grevillea (Mor et al. 1983; Lukaszewska et al.
1994; Taverner et al. 1999; Setyadjit et al. 2004), while it promoted flower senescence in
carnations (Woodson and Brandt 1991). The effects of cytokinins on plant development are
often dependent on the presence or the absence of other PGRs (Bernier 1988). For example,
GA enhanced the stimulatory effects of BA on inducing floral transition in Dendrobium
hybrids (Goh 1979; Bernier 1988). GA alone has been shown to promote flower opening in
Ipomoea nil and iris (Iris pseudacorus) (Raab and Koning 1987; Celikel and van Doorn
1995), and prolong longevity in alstromeria and carnation (Saks and van Staden 1993; Jordi
et al. 1995). However, GA was shown to have no effects on longevity of Grevillea and even
increased flower abscission (Setyadjit et al. 2006).
Other chemicals that had been shown to control flower opening include
aminooxyacetic acid (AOA), an inhibitor of ethylene synthesis (Rattanawisalanon et al.
2003). Treatment of Dendrobium ‘Jew Yuay Tew’ inflorescences with AOA suppressed bud
drop, promoted bud opening, and delayed flower senescence (Rattanawisalanon et al. 2003).
Longevity of Dendrobium ‘Heang Beauty’ flowers was also pro-longed upon treatment with
AOA (Chandran et al. 2006). It was proposed that AOA could act as an anti-microbial agent,
which inhibited bacterial growth and hence, allowing continuous uptake of water and sugars
by the flowers (Rattanawisalanon et al. 2003; Chandran et al. 2006).
2.5.3 Flowering and the senescence programme
There are suggestions that flowering might be regulated by a senescence programme
of the plant and/ or floral organs (Rubinstein 2000; O’Donoghue et al. 2002; Wagstaff et al.
2003). Senescence refers to the terminal phase in the development of leaves and flowers and
23
is often accompanied by events such as protein remobilization, increased proteinase
activities, DNA laddering, membrane degradation, cell wall alterations and nuclear
shrinkage, all of which are also characteristics of programmed cell death (PCD) (Rubinstein
2000; Wagstaff et al. 2003).
In Ipomoea, dynamic structural changes such as cell enlargement, modification of cell
shape and reduction in cell wall thickness occurred in the inner epidermal cells even before
flower opening (Phillips and Kende 1980). In sandersonia, intercellular air spaces and
increasingly disorganized packing of parenchyma cells also occurred prior to flower opening
(O’Donoghue et al. 2002). In alstromeria, DNA laddering, nuclear shrinkage and increase in
expression of cysteine protease all commenced as early as two days before flower opening
(Wagstaff et al. 2003). The occurrences of indicators of PCD before flower opening imply
that flower senescence is a continuum from a senescence programme that had already started,
and suggest that flower opening might somehow be a consequence of this senescence
programme (O’Donoghue et al. 2002; Wagstaff et al. 2003).
2.6 Orchids
Orchids originated from lily-like ancestors which have either evolved into orchids or become
extinct (Seidenfaden and Wood 1992). The classification of orchids is as such: superorder
Lilianae, order Orchidales, and family Orchidaceae (Seidenfaden and Wood 1992). The
column and the lip (or labellum) are the hallmarks of the orchids (Teoh 2005). Due to their
vividly coloured lips that are often embellished with crests, hair, ribs and other
protuberances, orchids are very appealing and are among the highly demanded cut flowers,
24
appreciated for their beauty and fragrance (Hew and Yong 2004; Teoh 2005; Chandran et al.
2006).
2.6.1 Dendrobium crumenatum
Dendrobium crumenatum (Swartz), also known as the pigeon orchid, is a common
native epiphytic orchid species of South-east Asia, naturally occurring in Singapore and
Malaysia (Fig. 1). It exhibits an interesting diversion of the normal flowering process: upon
transition of the meristem from a vegetative to a reproductive phase, floral buds develop to a
certain stage and then become ‘dormant’. These floral buds resume growth and development
after cold-induction, such as after a heavy rainfall, and culminating into the opening of the
flowers exactly nine days after (Holttum 1953; Corner 1988). Full flower opening is achieved
before dawn, and the white flowers are small (40 mm width) with yellow ridges running from
the centre of the lip to its base, providing the only colour in the flowers (Fig. 1E insets). The
flowers are short-lived, lasting only for a day before the onset of senescence (Tan and Hew
1993). The synchronized flowering of the pigeon orchid is an impressive sight, and the
cultivation of the orchid along roadside trees for aesthetic reasons has been attempted by the
National Parks Board of Singapore (Boo et al. 2006).
25
Fig. 1. Dendrobium crumenatum (pigeon orchid). An inflorescence
stalk bearing floral buds of day 6 (A), day 9 (B) and day 10, i.e. fully
opened flowers (C), after cold induction. (D) Synchronous flowering in
nature.
26
Chapter 3. Materials and Methods
3.1 Plant material
Plants of Dendrobium crumenatum (Swartz) were maintained under cool and partially shaded
conditions (PAR ranged from 100 – 250 µmol m-2 s-1; average air temperature ranged from
25 – 33°C) in a planthouse of the Department of Biological Sciences, National University of
Singapore. Plants were watered daily, and fertilized weekly with a foliar fertilizer (N:P:K =
18:36:18). Pots of D. crumenatum with inducible inflorescences carrying dormant floral buds
were acclimatized at 30°C for 24 h in temperature-controlled growth chambers. They were
then subjected to a cold induction at 20°C for 24 h. Growth chambers were maintained on a
12 h day/ 12 h night cycle and illumination was provided by fluorescent tubes (PAR ranged
from 10 – 20 μmol m-2 s-1). Plants of D. crumenatum exhibit crassulacean acid metabolism,
demonstrating different carbon dioxide exchange patterns during different times of the day.
Thus, all plants were moved into the growth chambers at 1600 h, to minimize the effects of
any possible temporal variations in the plant physiology. Plants were also watered daily to
minimize dehydration stress. Floral buds or flowers were selected according to their age and
features (Table 1, Fig. 5). For cell wall composition, cell wall enzyme activities and
molecular studies, sepals and petals of the harvested floral buds and flowers were separated
(Fig. 2) and stored at -80°C until use. Fresh samples were used for all other analyses.
3.2 Physical parameters analyses
Freshly harvested sepals and petals from each flower were weighed to obtain fresh weight,
then wrapped in aluminium foil and left to dry in an 80°C oven for 1 week for dry weight
27
Table 1. Stages of floral bud development in D. crumenatum. Timing of events is reported in
relation to the time during which floral buds were subjected to cold induction (denoted as day 0).
Features
Exposure of dormant floral buds to cold induction at 20°C for 24 h.
Time
(days after
induction)
0
Green bud (ca. 1 cm long) with reddish brown tinges along ventral side,
elongation of mentum, mentum reddish brown.
4
Light green bud (ca. 2.5 cm long), reddish brown tinges only at beginning
and tip of mentum, further elongation of mentum, length of mentum almost
half of length of whole bud.
7
White bud (ca. 3 cm long), no splitting of sepals, elongated mentum
pointing downwards away from tip of bud.
9
Full flower opening, sepals and petals fully expanded, lip fully protruded
with visible yellow ridges running down from midlobe to foot of column.
10
Sepals and petals shrivelled and brownish.
12
28
Fig. 2. Separation of floral parts in D. crumenatum floral buds and flowers. (A) Day 7
floral bud after dissection, (B) sepals of day 7 floral bud, (C) petals of day 7 floral
bud, (D) column of day 7 floral bud, (E) opened flower on day 10, (F) sepals of
opened flower, (G) petals of opened flower, (H) column of opened flowers. Scale bar
= 1 cm.
29
(amount of dry matter) determination. Water content was obtained by subtracting the amount
of dry matter from total fresh weight.
3.3 Microscopy
Segments of approximately 1 cm × 0.5 cm were cut about halfway from the tips of freshly
harvested sepals or petals, to include the central vein. The fresh tissues were fixed in 96 %
ethanol : acetic acid : formalin : water (10 : 1 : 2: 7 by volume) for 24 h. Fixed samples were
then subjected to dehydration through a graded tertiary-butanol (TBA) series (water : 95 %
ethanol : TBA) as follows: Grade 1 (4 parts of water : 5 parts of 95 % ethanol : 1 part of
TBA), 4 h at room temperature (22°C – 24°C) ; Grade 2 (3 : 5 : 2), 4 h at room temperature;
Grade 3 (1.5 : 5 : 3.5), 24 h at room temperature; Grade 4 (0 : 5 : 5), 4 h at room temperature;
Grade 5 (0 : 2.5 100 % ethanol : 7.5) with Orange-G (BDH) dye, 4 h at room temperature,
repeated once for 24 h; Grade 6 (0 : 0 : 10), 4 h at 40°C, repeated once. Dehydrated samples
were infiltrated with melted paraffin wax over 4 days at 60°C, then embedded in paraffin
wax. Embedded tissues were sectioned with a microtome to a thickness of 10 µm and
sections were mounted onto albumin-coated slides. The slides were dewaxed by Histoclear 1,
2 and 3 for 3 min each, and were hydrated through a series of decreasing concentration of
ethanol: 100 %, 95 %, 90 %, 70 %, 50 % and 0 % for 2 min each. The hydrated sections were
finally stained with toluidine blue (0.1 % in 0.1 M sodium phosphate buffer, pH 5.5) and
examined using light microscopy. The thickness of sepals or petals was obtained by
measuring the thickness of the area between the middle two vascular bundles of each section.
30
3.4 Electrolyte leakage
Electrolyte leakage was determined by the method described by Tuna et al. (2007) with some
modifications. Twenty discs (5 mm diameter) from freshly harvested sepals or petals were
obtained using a cork borer, and were equilibrated in 20 ml of water at room temperature for
1 h. Subsequently, electrical conductivity of the solution (C0) was measured using a
conductance meter (WTW Cond 315i/SET). Samples were boiled for 30 min, and electrical
conductivity of the solutions (C1) were measured after they cooled to room temperature. The
percentage of electrolyte leakage of sepals/ petals was calculated as follows:
Electrolyte leakage (%) = (C0/ C1) × 100
Membrane stability index (MSI) of sepals/ petals was calculated as follows:
MSI = [1 - (C0/ C1)] × 100
3.5 Cell wall compositional analyses
3.5.1 Preparation of EIR
Cell wall materials of sepals or petals were prepared by using a modified method described
by Huber (1992). Frozen tissues were homogenized in 95 % ethanol at 4°C and then chilled
at -20°C for 24 h. The homogenates were centrifuged at 8,000 g for 10 min at 4°C. Trisbuffered phenol (5 ml per g FW of sepal/ petal tissues) was added to the residues and allowed
to incubate at room temperature for 45 min. The suspensions were centrifuged as described
above, and the residues were re-suspended in 80 % ethanol at -20°C for 2 h, followed by
centrifugation. The remaining residues were washed once with 80 % ethanol, followed by 80
% acetone, and then chloroform : methanol (1 : 1) mixture. All organic washings were
conducted at room temperature. The final residues were recovered by filtration, washed
31
thrice with acetone until total whitening, yielding the crude cell wall material (ethanolinsoluble residue, EIR). The EIR was air-dried, weighed, and then stored at -80°C until use.
The amount of EIR was expressed as mg per gram fresh weight and mg per floral part (sepal
or petal).
3.5.2 Cellulose content
Cellulose was extracted from 5 mg of EIR following the procedure described by Updegraff
(1969). Initial hydrolysis was carried out in 5 ml of acetic-nitric acid reagent (10 volumes of
80 % acetic acid : 1 volume of 16N nitric acid) at 100°C for 30 min. Samples were
centrifuged at 4,000 g for 5 min at room temperature, and the supernatants discarded. The
remaining residues were washed with 10 ml of water, centrifuged as described above, and the
supernatants were discarded. The remaining residues were hydrolysed in 10 ml of 67 %
sulphuric acid for 1 h at room temperature. The resultant solution was diluted 10 times with
water and was analysed for cellulose content.
Cellulose contents were assayed by the anthrone method (Scott and Melvin 1953).
Four ml of water were added to 1 ml of sample in an ice-water bath and then 10 ml of chilled
anthrone reagent (Sigma) were added. The sample was mixed, and then incubated at 100°C
for 15 min. The reaction was stopped by immediately returning the sample to an ice-water
bath. The bluish-green colour of the mixture was allowed to develop at room temperature for
10 min and the absorbance of the mixture at 620 nm was measured. Cellulose (0 – 0.2 mg ml1
, Sigma) was used as the standard in the assays. Cellulose concentration was expressed as
mg per gram fresh weight and μg per floral part (sepal or petal).
32
3.5.3 Total pectin content
Total pectins in EIR (5 mg) were extracted as described by Ahmed and Labavitch (1977).
Two ml of chilled, concentrated sulphuric acid were added to the samples and allowed to
incubate for 5 min. This was followed by the addition of 0.5 ml of water to the mixture and
incubated for 5 min. Another 0.5 ml of water were added until dissolution of the materials
was completed. The whole extraction procedure was conducted in an ice-water bath with
constant gentle swirling. The hydrolysed sample was made up to a total of 10 ml with water
and the resultant solution was analysed for total pectin content.
Pectin contents were expressed as uronic acid equivalents and were determined by the
m-hydroxydiphenyl method (Blumenkrantz and Asboe-Hansen 1973). Six ml of chilled
0.0125 M sodium tetraborate-sulfuric acid solution (Sigma) were added to 1 ml of sample on
an ice-water bath. The mixture was mixed well, and then incubated at 100°C for 6 min. The
reaction was terminated by immediately returning the mixture to an ice-water bath. This was
followed by the addition of 0.1 ml of m-hydroxydiphenyl (0.15 % w/v, Fluka) to the sample.
The pink colour of the mixture was allowed to develop for 15 min at room temperature and
the absorbance at 520 nm was measured. Galacturonic acid (0 – 0.2 mg ml-1, Fluka) was
used as the standard in the assays. Total pectin concentration was expressed mg per gram
fresh weight and μg per floral part (sepal or petal).
3.5.4 Soluble pectin and hemicellulose contents
Soluble pectins and hemicelluloses were sequentially extracted from EIR (Fig. 3). The
extractions were carried out in water (water-soluble pectins), 50 mM sodium acetate buffer
(pH 6.0) containing 50 mM CDTA (CDTA-soluble pectins), 50 mM Na2CO3 containing 20
mM NaBH4 (Na2CO3-soluble pectins), and finally 6 N NaOH containing 0.13 mM NaBH4
33
Fig. 3. Sequential extraction of pectins and hemicelluloses from EIR of D. crumenatum
floral buds.
34
(hemicelluloses). The proportion of EIR to extractant was 1 mg : 0.5 ml. Each extraction was
conducted at room temperature for 2 h with constant shaking, and centrifuged at 10,000 g for
15 min at 4°C (O’Donoghue et al. 2002; Deng et al. 2005).
The various supernatants were analysed for their respective contents. Hemicellulose
contents were assayed by the anthrone method (Scott and Melvin 1953) described above and
pectin contents were assayed by the the m-hydroxydiphenyl method (Blumenkrantz and
Asboe-Hansen 1973) described above. Hemicellulose and soluble pectin concentrations were
expressed as mg per gram fresh weight and μg per floral part (sepal or petal).
3.6 Cell wall enzyme analyses
3.6.1 Enzyme extraction
Sepals and petals were ground to fine powder in liquid nitrogen with a mortar and
pestle. The extraction buffer was then added and the materials were homogenized further. All
enzyme extractions were carried out at 4°C. The homogenates were centrifuged at 12,000 g
for 20 min at 4°C, and the supernatants were used to determine enzymatic activities.
For cellulase (EC 3.2.1.4) and polygalacturonase (PG; EC 3.2.1.15), two extraction
methods (Method A and B) were attempted on both fresh and frozen plant materials, as
summarized in Table 2. Method A was a 1-step procedure extracting salt-soluble cellulase
and PG with 20 mM citrate buffer (pH 5.1) containing 1 M NaCl, or 20 mM sodium
phosphate buffer (pH 6.1 or pH 7) containing 1 M NaCl (Panavas et al. 1998). The
extractions were conducted over 30 min. Method B was a 4-step procedure extracting both
buffer-soluble and salt-soluble cellulase and PG (Ferrari and Arnison 1974; Huberman et al.
35
Table 2. Extraction procedures of cellulase and PG from floral buds of D. crumenatum. Method
A was a 1-step procedure (Panavas et al. 1998), while Method B was a 4-step procedure (Ferrari
and Arnison 1974; Huberman et al. 1975). The type of supernatants collected from each step of
extraction is included.
Method
Extraction procedure
Supernatant
A
1. Tissue homogenized in either
- 20 mM citrate buffer (pH 5.1) + 1 M NaCl
or
- 20 mM sodium phosphate buffer (pH 6.1) + 1 M
NaCl
or
- 20 mM sodium phosphate buffer (pH 7) + 1 M
NaCl
Salt-soluble
cellulase and
PG
B
1. Tissue homogenized in either
- 20 mM citrate buffer (pH 5.1)
or
- 20 mM sodium phosphate buffer (pH 6.1)
or
- 20 mM sodium phosphate buffer (pH 7)
Buffersoluble
cellulase and
PG/
2. Residue washed with respective homogenizing buffer.
3. Residue washed with respective homogenizing buffer.
4. Residue resuspended in respective homogenizing buffer +
1 M NaCl for 30 min.
Salt-soluble
cellulase and
PG
36
1975). The materials were homogenized for 30 min in 20 mM citrate buffer (pH 5.1) or 20
mM sodium phosphate buffer (pH 6.1 or pH 7). The homogenates were centrifuged to obtain
the supernatants (buffer-soluble enzyme) and the remaining residues were washed twice with
the respective homogenizing buffers and, after centrifugation, the supernatants were
discarded. The final remaining residues were resuspended in the respective buffers with 1 M
NaCl for 30 min, and centrifuged to obtain the supernatants (salt-soluble enzyme) (Ferrari
and Arnison 1974; Huberman et al. 1975). The ratio of plant material to extractant ranged
between 1g : 10 ml to 1g : 1 ml. Aliquots of the various supernatants were desalted on
Sephadex G-25 columns (1 × 10 cm). All extracts (desalted and crude) were tested for
activities of both cellulase and PG.
For the extractions of pectin methylesterase (PME; EC 3.1.1.11), β-galactosidase (βgal; EC 3.2.1.23), β-glucosidase (β-glu; EC 3.2.1.21), β-mannosidase (β –man; EC 3.2.1.25)
and β-xylosidase (β –xyl; EC 3.2.1.37), frozen plant materials were used. Plant materials
were homogenized in 20 mM sodium phosphate (pH 7.5) containing 1.5 M NaCl for 30 min.
The ratio of plant material to extractant was 1 g : 10 ml. The supernatants were recovered by
centrifugation and the crude extracts were used for the various enzymatic assays.
3.6.2 Cellulase assay
A gel diffusion assay (Wood et al. 1988) was adopted to screen for cellulase activity. Gels,
consisting of 1.7 % agarose and 0.5 % carboxymethylcellulose (CMC; medium viscosity,
Sigma) in 0.2 M citrate buffer (pH 5.1) or 0.2 M sodium phosphate buffer (pH 6.1 or 7), were
prepared in Petri-dishes (diameter = 9 cm) to a thickness of approximately 4 mm. Wells of 5
mm diameter were cut using a cork borer, and the cellulase extracts (25 μl) were pipetted into
the wells. The Petri-dishes were sealed with parafilm and were incubated at room
37
temperature or 37°C for 2 h or 24 h. The gels were then stained by covering the plates with 1
mg ml-1congo red solution (BDH) for 15 min, followed by 1 M NaCl for 10 min. The NaCl
wash was repeated twice. Stained plates were preserved for later observations by flooding
with 5 % acetic acid. Commercial cellulase (0.5 mg ml-1, Yakult Honsha) and boiled enzyme
extracts were included as experimental controls.
Cellulase activity was also assayed by spectrophotometric methods adapted from
Ghose (1987). Using CMC as the enzyme substrate, the reaction mixture contained 0.4 ml of
0.05 % CMC, 0.2 ml of 0.1 M sodium acetate buffer (pH 5) and 400 μl of enzyme extract.
Using filter paper as the enzyme substrate, the reaction mixture contained a filter paper strip
(1 cm × 1 cm, Whatman No. 1), 0.6 ml of 0.1 M sodium acetate buffer (pH 5) and 400 μl of
enzyme extract. Incubation was carried out at 37°C for 1 h or 24 h. In control tubes, boiled
enzyme extracts were used. The amounts of reducing groups released from CMC and filter
paper strips were determined by the dinitrosalicylic acid (DNS) and glucose oxidase/оdianisidine (GOD) methods (Ghose 1987). In the DNS method, 2 ml of DNS reagent,
consisting of 0.02 M DNS (Sigma), 2 N NaOH and 1 M Rochelle salt (Sigma), were added to
the reaction mixture, and then heated in a boiling water bath for 5 min. The mixtures were
immediately placed in a ice-water bath for 5 min after the heat treatment, and 9 ml of water
were added to each tube. The absorbance at 540 nm was measured. In the GOD method, 2 ml
of assay reagent (glucose oxidase/о-dianisidine, Sigma) were added to 0.5 ml of reaction
mixture, and incubated at 37°C for 30 min. The reactions were stopped by the addition of 2
ml of 12 N H2SO4. The absorbance at 540 nm was measured. Glucose (0 – 2 mg ml-1 for the
DNS assay, 0 – 0.16 mg ml-1 for the GOD assay) was used as the standard in these assays.
One unit of cellulase activity represented 1 μmol of reducing groups liberated per h.
38
Cellulase activity was expressed as units per mg protein, units per floral part (sepal or petal)
and units per g fresh weight.
3.6.3 Polygalacturonase assay
Polygalacturonase (PG) activity was determined by an adapted method described by AbuBakr et al. (2003). The reaction mixture, containing 0.4 ml of 0.1 % polygalacturonic acid
solution (Fluka), 0.2 ml of 0.1 M sodium acetate buffer (pH 5) and 400 μl of enzyme extract,
was incubated at 37°C for 1 h or 24 h. In control tubes, boiled enzyme extracts were used.
The amount of reducing groups released from polygalacturonic acid was determined by the
DNS and GOD methods as described above. Galacturonic acid (0 – 2 mg ml-1 for the DNS
assay, 0 – 0.16 mg ml-1 for the GOD assay) was used as the standard in these assays. One unit
of PG activity represented 1 μmol of reducing groups liberated per h. PG activity was
expressed as units per mg protein, units per floral part (sepal or petal) and units per gram
fresh weight.
3.6.4 Pectin methylesterase assay
Pectin methylesterase (PME) activity was assayed by a continuous spectrophotometric
method as described by Hagerman and Austin (1986) with some modifications. The reaction
mixture contained 2 ml of 0.5 % citrus pectin solution (degree of esterification = ~60 %,
Sigma), 0.15 ml of 0.01 % bromothymol blue (Sigma) solution in 3 mM potassium
phosphate buffer (pH 7.5), 0.75 ml of water and 100 μl of enzyme extract. The pHs of the
solutions were adjusted to pH 7.5 each time before use. After adding the enzyme extract, the
reaction mixture was mixed well and allowed to stabilize for 1 min at room temperature
before measuring the absorbance at 620 nm. The absorbance was again determined after 21
min of incubation at room temperature. The difference in absorbance was the measure of
39
PME activity and was calibrated against a galacturonic acid standard curve (0 – 0.17 mg ml1
). One unit of PME activity represented 1 µmol of methylester liberated per h. Boiled
enzyme extracts were included as controls. PME activity was expressed as units per mg
protein, units per floral part (sepal or petal) and units per gram fresh weight.
3.6.5 Glycosidases assay
β-galactosidase (β-gal) , β-glucosidase (β-glu), β-mannosidase (β –man) and β-xylosidase (β
–xyl) activities were assayed by an adapted method described by Chin et al. (1999). The
reaction mixtures consisted of 0.5 ml 5 mM p-nitrophenyl derivatives of β-Dgalactopyranoside, β-D-glucopyranoside, β-D-mannopyronoside or β-D-xylopyranoside
(Sigma) as substrate, 50 mM sodium acetate buffer (pH 4.5) and 50 μl of enzyme extract in a
total volume of 2 ml. After incubation at 37°C for 30 min, the reactions were stopped by
adding 1 ml of 0.2 M Na2CO3, and the amount of p-nitrophenol formed was determined
spectrophotometrically at 415 nm. One unit of glycosidase activity represented 1 μmol of pnitrophenol released per h. Boiled enzyme extracts were included as controls. Glycosidase
activity was expressed as units per mg protein, units per floral part (sepal or petal) and units
per gram fresh weight.
3.6.6 Soluble protein content
Soluble protein concentration was determined by the Bradford method (1976) with
bovine serum albumin (0 – 0.9 mg ml-1) as the standard. Three ml of 20 % (v/v) Bio-Rad
reagent were added to 60 μl of extract. The reaction mixture was incubated at room
temperature for 20 min. The absorbance at 595 nm was then measured. Soluble protein
concentration was expressed as mg per gram fresh weight and μg per floral part (sepal or
petal).
40
3.7 Gene expression profiling
All reagents, centrifuge tubes and pipette tips were autoclaved at 121°C for 60 min and
mortars/ pestles were baked at 180°C for at least 16 h to ensure that they were RNase free.
3.7.1 Total RNA isolation
Total RNA was extracted from frozen sepals and petals. Samples were ground in a mortar
and a pestle in liquid nitrogen and Trizol reagent (Invitrogen) was added (1 ml per 0.1 g fresh
weight of plant materials). The samples were further ground into fine powder in liquid
nitrogen and allowed to thaw at room temperature. The samples were then centrifuged at
12,000 g for 10 min at 4°C. The resultant supernatant was transferred to a new centrifuge
tube and chloroform (0.2 ml per 1 ml Trizol) was added. This was followed by vortexing at
high speed for 15 s, and incubation at room temperature for 4 min. The samples were
centrifuged at 14,000 g for 15 min at 4°C, and the colourless aqueous phases were transferred
to new centrifuge tubes. The chloroform extraction was repeated once.
To the final aqueous phase, isopropanol and 0.8 M sodium citrate/1.2 M NaCl were
added (0.5 ml each per 1 ml of aqueous phase), and mixed by gentle inversion. The samples
were incubated on ice for 20 min, and then centrifuged at 14,000 g for 20 min at 4°C. The
supernatants were discarded, and pre-cooled 75 % ethanol (same volume at Trizol) was
added. Samples were vortexed briefly then centrifuged again at 7800 g for 10 min at 4°C.
The supernatants were discarded and the resulting pellets were dried under vacuum for 5
min. The dried pellets were solubilized in RNase-free water and an equal volume of 8 M
lithium chloride was added. The samples were mixed, and then incubated at -80°C overnight.
The frozen samples were allowed to thaw on ice, and then centrifuged at 12,000 g for
20 min at 4°C. The supernatants were discarded and the remaining residues were washed
41
with pre-cooled 75 % ethanol (same volume as Trizol). Samples were vortexed briefly, and
then centrifuged at 7800 g for 10 min at 4°C. The supernatants were discarded and the
resulting pellets were dried under vacuum for 5 min. The dried pellets were solubilized in
RNase-free water and then stored at -80°C.
3.7.2 Estimation of RNA quality and quantity
The amount of RNA (ng/μl) was quantified spectrophotometrically. Purity of RNA
was determined by the absorbance ratios A260/280 and A260/230, which are measures of
contamination by proteins and polyphenols/ carbohydrates respectively. The integrity of
RNA was also verified by analyzing approximately 1 μg RNA sample on 0.8 % agarose gels.
3.7.3 Reverse transcription
First-strand cDNA synthesis was conducted using the SuperScript First-Strand Synthesis
System for RT-PCR kit (Invitrogen). Approximately 100 ng of RNA were added to 1 μl of
10 mM dNTP, 1 μl of oligodT (0.5 ng/μl) and DEPC-treated water to a total volume of 10 μl.
The samples were incubated at 65°C for 5 min, then incubated at 4°C for 2 min. This was
followed by the addition of 2 μl of 10x RT buffer, 4 μl of 25 mM MgCl2, 2 μl of 0.1 M DTT
and 1 μl of RNaseOUT Recombincat RNase Inhibitor. The mixtures were mixed, and then
incubated at 42°C for 2 min. One μl (50 units) of SuperScript II RT was then added to each
sample, mixed, and incubated at 42°C for 50 min. The reactions were terminated at 70°C for
15 min, and then chilled at 4°C. The final cDNA obtained was stored at -20°C.
42
3.7.4 PCR amplification
The primers used for PCR are as follows: β-TUB (β-tubF, 5’-CGTAAGGAAGCTGAGAACTGTGATTGC-3’, and β-tubR, 5’- GCAAGAAAGCTTTACGCCTGAACATAG-3’); β-GAL
(β-galF, 5’- CCTATGTGTTCTGGAACGGGC-3’, and β-galR, 5’-CATCTTCCTTGCACATGACCCATGG-3’); PME (PMEF, 5’-GCACCGTCGACTTCATCTTC-3’, and PMER, 5’GGCATATACCCCTCAGG-3’). PCR amplification was performed using the GoTAQ Flexi
DNA Polymerase (Promega). Gradient PCR was performed at various temperatures to
determine the optimum annealing temperatures for the respective pairs of primers. PCR
amplification was also conducted over 24, 26, 28, 30 and 32 cycles to determine the plateau
phase for each gene. The final optimized PCR conditions were as follows: 95°C for 5 min, 30
cycles of 94°C for 30 s, 52°C for 1 min, 72°C for 1 min and 72°C for 5 min. The amplified
samples were analysed by gel electrophoresis on 1.2 % agarose gel containing GelRed stain
(8 μl per 100 ml agarose, Biotium Inc.).
3.8 Controlling time of flower opening in D. crumenatum
3.8.1 General setup
Inflorescences of D. crumenatum bearing mature floral buds were harvested, and each floral
bud was then cut such that it included the stalk of the bud and a short section of the
inflorescence stem. Individual buds were placed in 2 ml centrifuge tubes containing the test
solution. The centrifuge tubes holding the floral buds were then placed into GA7 containers,
and were sealed with parafilm. Holes were made in the parafilm to facilitate transpiration,
and to prevent condensation in the GA7 containers. The GA7 containers were then placed on
a culture rack under a 16 h light/ 8 h dark cycle (PAR = 32 μmol m-2 s-1) at room
43
temperature. The floral buds were examined for signs of flower opening over the next few
days.
3.8.2 Treatment
The excised floral buds were treated with different concentrations of benzyladenine (BA,
Sigma) (10-6 M, 10-8 M, 10-10 M), gibberellic acid (GA3, Sigma) (10-6 M, 10-8 M, 10-10 M), a
combination of BA/ GA3 (10-6 M, 10-8 M, 10-10 M) and aminooxyacetic acid (AOA, Sigma)
(10-6 M, 10-8 M, in the presence and absence of 0.1 M glucose). Water was included as a
control in the study.
3.9 Statistical analysis
The results were presented as mean ± standard error. The results were analysed for statistical
significance by multifactor ANOVA. Where ANOVA detected significant differences,
Fisher’s least significance difference (LSD) test was used at a 5 % level of significance.
44
Chapter 4. Results
4.1 Growth changes
Sepals and petals of D. crumenatum demonstrated similar patterns of weight changes (Fig.
4). Total fresh weight (Fig. 4A) increased during floral bud development (Phase I); it
remained constant during flower opening (Phase II) and decreased upon flower senescence
(Phase III). The increases in water content during floral bud development (Phase I) followed
an exponential pattern, with a dramatic increase occurring during the late stages of floral bud
development on day 9 (Fig. 4B), whereas the increases in dry matter content during floral
bud development were relatively constant (Fig. 4C). During flower opening (Phase II), while
no changes were observed for water content, dry matter content decreased slightly (Fig. 4B,
C). Upon flower senescence (Phase III), dry matter content remained constant (Fig. 4C),
while water content decreased significantly (P < 0.05), accounting for the significant
decreases (P < 0.05) in total fresh weight and FW : DW ratio (Fig. 4A, B, C).
4.2 Changes in anatomy during development
Cross-sections of sepals and petals from young D. crumenatum floral buds (day 4) showed
round parenchyma cells packed tightly and neatly between the upper and lower epidermes
(Fig. 5A). During floral bud development (Phase I), the parenchyma cells were less tightly
packed together, exhibiting some intercellular spaces (Fig. 5B, C). This layer became
progressively disorganized during flower opening (Phase II), with larger intercellular spaces
and membrane disintegration becoming apparent (Fig. 5D). During flower senescence (Phase
45
Phase
I
II
III
Fig. 4. Characteristics of weight changes of sepals and petals of D. crumenatum. (A)
Fresh weight, (B) Water content, (C) Dry matter content and FW : DW ratio. Phases of
development – I: floral bud development, II: flower opening, III: flower senescence.
Each value is the mean ± SE (n = 5), and values with different alphabet(s) (sepals: a – d;
petals: a’ – d’) are significantly different according to Fisher’s LSD test (P < 0.05).
46
Stage
Sepal
Petal
I
Phase
II
III
Fig. 5. Changes in anatomical features of D. crumenatum during flower
development. Developmental stages of floral buds/ flowers after cold induction:
A, day 4; B, day 7; C, day 9; D, day 10; E, day 12 floral buds/ flowers. Features
of development were as described in Table 1. Phases of development – I: floral
bud development, II: flower opening, III: flower senescence. Transverse sections
of sepals and petals were stained with toluidine blue. Arrows point to areas with
parenchyma cell disruption and /or intercellular spaces. Anatomical studies were
repeated four times on different floral buds or flowers.
47
III) on day 12, both sepals and petals showed almost complete absence of intact parenchyma
cells, although the epidermal layers remained rather intact (Fig. 5E).
The development of D. crumenatum floral buds was accompanied by marked changes
in cell size (Fig. 6). The thickness of sepals and petals increased significantly (P < 0.05)
during floral bud development (Phase I), reaching a maximum on day 9; it subsequently
decreased during flower opening (Phase II) and flower senescence (Phase III) (Fig. 6A). The
pattern of changes in sepals/ petal thickness corresponded with the changes in individual cell
length. The lengths of parenchyma cells in sepals and petals were maximum during late floral
bud development (day 9); it then decreased significantly (P < 0.05) upon full flower opening
and senescence (Fig. 6C). Cell width of parenchyma cells, on the other hand, increased
throughout development (Fig. 6B).
4.3 Cell wall composition
4.3.1 EIR
Sepals and petals demonstrated similar patterns of changes in cellulose content throughout
development. On a per floral part (sepal or petal) basis, the amount of EIR increased during
floral bud development (Phase I) and flower opening (Phase II), from approximately 0.2 mg
(floral part)-1 to 0.8 mg (floral part)-1 (Fig. 7A). A significant loss in EIR was observed,
decreasing to approximately 0.4 mg (floral part)-1, during flower senescence (Phase III).
On a per gram fresh weight basis, EIR content decreased significantly (P < 0.05)
during floral bud development (Phase I); it then remained at low and constant levels
throughout flower opening (Phase II) and flower senescence (Phase III) (Fig. 8A).
48
I
Phase
II
III
Fig. 6. Anatomical changes during D. crumenatum floral bud development. (A) Sepal
and petal thickness measured between two middle vascular bundles, (B) width of
parenchyma cells, (C) height of parenchyma cells. Phases of development – I: floral bud
development, II: flower opening, III: flower senescence. Each value is mean ± SE (n =
5), and values with different alphabet(s) (sepals: a – c; petals: a’ – c’) are significantly
different according to Fisher’s LSD test (P < 0.05).
49
I
Phase
II
III
I
I
Phase
II
Phase
II
III
III
Fig. 7. D. crumenatum sepal and petal cell wall components, expressed on a per floral part (sepal/
petal) basis, at each developmental stage. (A) EIR, (B) cellulose, (C hemicelluloses, (D pectins, (E)
water-soluble pectins, (F) CDTA-soluble pectins, (G) Na2CO3-soluble pectins. Phases of
development – I: floral bud development, II: flower opening, III: flower senescence. Each value is
mean ± SE (n = 5) and values with different alphabet(s) (sepals: a – d; petals: a’ – d’) are
significantly different according to Fisher’s LSD test (P < 0.05).
50
I
Phase
II
III
I
I
Phase
II
Phase
II
III
III
Fig. 8. D. crumenatum sepal and petal cell wall components, expressed on a per gram fresh weight
(gFW-1) basis, at each developmental stage. (A) EIR, (B) cellulose, (C) hemicellulose, (D) pectin,
(E) water-soluble pectin, (F) CDTA-soluble pectin, (G) Na2CO3-soluble pectin. Phases of
development – I: floral bud development, II: flower opening, III: flower senescence. Each value is
mean ± SE (n = 5) and values with different alphabet(s) (sepals: a – c; petals: a’ – c’) are
significantly different according to Fisher’s LSD test (P < 0.05).
51
4.3.2 Cellulose
Sepals and petals demonstrated similar patterns of changes in cellulose content throughout
development. On a per floral part (sepal or petal) basis, cellulose content increased
significantly (P < 0.05) during floral bud development (Phase I) from approximately 5 μg
(floral part)-1 to 20 μg (floral part)-1; it continued to increase during flower opening (Phase II)
to approximately 30 μg (floral part)-1 (Fig. 7B). Upon flower senescence (Phase III),
cellulose content decreased significantly, reducing to approximately 18 μg (floral part)-1.
On a per gram fresh weight basis, a general increase in cellulose content was
observed during floral bud development (Phase I), flower opening (Phase II) and flower
senescence (Phase III) (Fig. 8B).
4.3.3 Hemicelluloses
Sepals and petals demonstrated similar patterns of changes in hemicellulose content
throughout development. On a per floral part (sepal or petal) basis, the amount of
hemicelluloses decreased constantly throughout floral bud development (Phase I), flower
opening (Phase II) and flower senescence (Phase III), decreasing from approximately 30 μg
(floral part)-1 on day 4 (Phase I) to 2 μg (floral part)-1 on day 12 (Phase III) (Fig. 7C).
On a per gram fresh weight basis, hemicellulose content also decreased constantly
throughout floral bud development till flower senescence (Fig. 8C).
4.3.4 Total pectins
Sepals and petals demonstrated similar patterns of changes in total pectin content throughout
development. On a per floral part basis, the amount of total pectins increased during floral
bud development (Phase I), increasing from approximately 30 μg (floral part)-1 on day 4 to
52
80 μg (floral part)-1 on day 9. No changes were observed in total pectin content during flower
opening (Phase II) and flower senescence (Phase III) (Fig. 7D).
On a per gram fresh weight basis, a decrease in total pectin content was observed
during early stages of floral bud development (Phase I); it remained constant till flower
opening (Phase II) and increased upon flower senescence (Phase III) (Fig. 8D).
4.3.5 Soluble pectins
Sepals and petals demonstrated similar patterns of changes in soluble pectin content
throughout development. On a per floral part basis, the quantities of water-soluble, CDTAsoluble and Na2CO3-soluble pectins increased during floral bud development (Phase I) (Fig.
7E – G). While the increases in CDTA-soluble and Na2CO3-soluble pectins during this
developmental period were relatively constant, that of water-soluble pectins was very drastic,
with an approximately three fold increase in content during the early stages of floral bud
development (day 7). The amount of CDTA-soluble pectins remained at high levels
throughout flower opening (Phase II) and flower senescence (Phase III). On the other hand,
the quantities of water-soluble and Na2CO3-soluble pectins decreased upon flower opening
and flower senescence respectively. When expressed as a percentage of total pectins, all
soluble pectins demonstrated an increase during floral bud development (Phase I), and
subsequently decreased during flower senescence (Phase III) (Table 3, 4). In sepals, watersoluble pectins increased from 5.5 % to 9.1 % during floral bud development, then decreased
to 4.6 % upon senescence; CDTA-soluble pectins increased from 13.5 % to 24.5 % during
floral bud development, then decreased slightly to 22.8 % upon senescence; Na2CO3-soluble
pectins increased from 36.9 % to 63.0 % during floral bud development, then decreased to
50.1 % upon senescence (Table 3). The corresponding changes in petals were 5.1 % to 6.7 %
53
Table 3. Total and soluble pectins in EIR derived from D. crumenatum sepals at various
developmental stages. Numbers in parentheses represent solubilized pectins as a percentage of
total pectins.
Stage of
development
(days after
induction)
Total
pectins
[μg (floral
part)-1]
4
25.2
1.4
(5.5)
3.4
7
70.5
6.4
(9.1)
14.2 (20.1)
42.1 (59.7)
9
90.5
6.6
(7.3)
22.2 (24.5)
57.0 (63.0)
II
10
95.4
4.9
(5.1)
21.2 (22.2)
55.9 (58.6)
III
12
90.6
4.2 (4.6)
20.7 (22.8)
45.4 (50.1)
Phase of
development
I
Soluble pectins [μg (floral part)-1]
Watersoluble
CDTAsoluble
(13.5)
Na2CO3soluble
9.3
(36.9)
54
Table 4. Total and soluble pectins in EIR derived from D. crumenatum petals at various
developmental stages. Numbers in parentheses represent solubilized pectins as a percentage of
total pectins.
Stage of
development
(days after
induction)
Total
pectins
[μg (floral
part)-1]
Watersoluble
CDTAsoluble
4
23.4
1.2 (5.1)
1.3 (5.6)
6.3
7
45.1
3.0 (6.7)
7.4 (16.4)
27.4 (60.8)
9
60.3
3.2 (5.1)
8.5 (14.1)
38.9 (64.5)
II
10
63.5
3.3 (5.2)
9.4 (14.8)
35.4 (55.7)
III
12
61.0
1.5 (2.5)
7.8 (12.8)
28.5 (46.7)
Phase of
development
I
Soluble pectins [μg (floral part)-1]
Na2CO3soluble
(26.9)
55
to 2.5 % for water-soluble pectins, 5.6 % to 16.4 % to 12.8 % for CDTA-soluble pectins, and
26.9 % to 64.5 % to 46.7 % for Na2CO3-soluble pectins (Table 4). Maximum percentages of
water-soluble, CDTA-soluble and Na2CO3-soluble pectins were all observed during the
intermediate (day 7) or late (day 9) stages of floral bud development (Phase I), indicating a
maximum level of pectin solubility during this developmental period.
On a per gram fresh weight basis, a constant decrease in water-soluble pectin content
was observed throughout floral bud development (Phase I), while decreases in contents of
CDTA-soluble and Na2CO3-soluble pectins were observed only during the later stages of
floral bud development (Fig. 8E – G). During flower opening (Phase II), decreases in
amounts of CDTA-soluble and Na2CO3-soluble pectins were also observed. Upon flower
senescence (Phase III), while content of water-soluble pectins remained at low and constant
levels, contents of CDTA-soluble and Na2CO3-soluble pectins increased (Fig. 8E – G).
4.4 Activities of cell wall-based enzymes
4.4.1 Soluble proteins
Sepals and petals demonstrated similar patterns of changes. On a per floral part basis,
soluble protein content increased steadily during floral bud development (Phase I) and flower
opening (Phase II) (Fig. 9A). Between young floral buds (day 4) and newly opened flowers
(day 10), soluble protein content increased by approximately two folds. A drastic and
significant (P < 0.05) decrease in soluble protein content of approximately eight folds was
observed during flower senescence (Phase III).
56
Phase
I
II
III
Fig. 9. D. crumenatum sepal and petal soluble protein content at each developmental
stage. (A) Expressed on a per floral part basis; (B) expressed on a per gram fresh weight
(gFW-1) basis. Phases of development – I: floral bud development, II: flower opening,
III: flower senescence. Each value is mean ± SE (n = 5) and values with different
alphabet(s) (sepals: a – e; petals: a’ – e’) are significantly different according to Fisher’s
LSD test (P < 0.05).
57
On a per gram fresh weight basis, soluble protein levels decreased during floral bud
development (Phase I), and remained at low and relatively constant levels throughout flower
opening (Phase II) and flower senescence (Phase III) (Fig. 9B).
4.4.2 Cellulase
Optimization of the cellulase gel diffusion assay using commercially available cellulase
demonstrated that the assay, when applied on acidic gel, provided the most sensitive results
(Fig. 10). Increasing contrast from the congo red staining was observed with increasing
acidity of the gel, aiding the measurement of the diffusion zone (Fig. 10A, B, C). The pH of
the gel only affected the clarity of the diffusion zones and did not affect cellulase activity
(Fig. 10D – R). Cellulase activity increased with higher incubation temperature and longer
incubation duration (Fig. 10D – R).
Due to the different pH values of the gel and cellulase solutions, a pH gradient would
have existed upon the diffusion of the enzyme into the gel. The comparison of commercially
available cellulase activity in such a setup was required as the extraction of cellulase from the
sepals and petals of D. crumenatum was conducted over various pH as well (refer to Section
3.6.1 under Materials and Methods). In the presence or absence of a pH gradient, the extent
of zone clearing at each gel pH was observed to be unaffected by the pH of the cellulase
solutions (Fig. 10D – R). Subsequently, the gel diffusion assay for D. crumenatum extracts
was conducted on pH 5.7 gel.
When extracted at pH 5.1, desalted extracts from fresh D. crumenatum sepals and
petals of all developmental stages did not exhibit any zone clearing. Zone clearance was
however observed using desalted extracts from fresh D. crumenatum column tissues
extracted at pH 5.1 (Fig. 11). Similar results were obtained with crude extracts from fresh
58
59
2h
22°C
24h
2h
37°C
24h
24 h
37°C
Fig. 10. Optimisation of cellulase gel diffusion assay using commercial cellulase (0.5 mg ml-1). Gels (1.7 %
agarose) contained 0.5 % CMC as substrates. Zone clearing for assays conducted at gel pH 5.7, pH 6.1 and pH
7.0, at 22°C for 2 h (D – F), 22°C for 24 h (G – H), 37°C for 2 h (J – L), 37°C for 24 h (A – C, M – O). Gels
without CMC were included as controls (P – R). Positioning of wells: 1) cellulase (pH 5.1); 2) boiled cellulase
(pH 5.1); 3) cellulase (pH 7.0); 4) boiled cellulase (pH 7.0).
2h
37°C
Fig. 11. Gel diffusion assay for cellulase. Assays were carried out at gel pH 5.7 at
37°C for a 24 h incubation period; 0.5% CMC as substrate. (A) Day 4, (B) Day 7, (C)
Day 9, (D) Day 10, (E) Day 12 after cold induction, (F) position of wells and desalted
samples extracted at pH 5.1. Positioning of wells: 1) sepals, 2) boiled sepals, 3) petals,
4) boiled petals, 5) columns, 6) boiled columns, 7) commercial cellulase (0.5 mg ml-1),
8) boiled commercial cellulase. Similar results were obtained from samples extracted
and assayed at pH 6.1 and 7. Lower concentrations of CMC (0.05% and 0.1%) also
showed similar results.
60
tissues, and when tissues were extracted at pH 6.1 and 7.0. No cellulase activities in all sepal
and petal extracts were observed in tests with lower concentrations of CMC (0.05 % and 0.1
%). All extracts from frozen tissues exhibited no cellulase activity.
Colorimetric analyses using crude or desalted extracts from fresh sepal or petal
tissues, regardless of extraction pH, demonstrated that the extracts did not contain
components to hydrolyze CMC and filter paper. Crude or desalted extracts from column
tissues, however, demonstrated cellulase activity. It was concluded that sepals and petals of
D. crumenatum had no or very insignificant levels of cellulase activity and no further studies
were conducted.
4.4.3 Polygalacturonase
Colorimetric analyses using crude or desalted extracts from fresh sepal or petal
tissues, regardless of extraction pH, did not contain components to hydrolyze
polygalacturonic acid. Desalted extracts from fresh or frozen D. crumenatum column tissues
however, demonstrated PG activity. It was concluded that sepals and petals of D.
crumenatum had no or very insignificant levels of PG activity and no further studies were
conducted.
4.4.4 Pectin methylesterase
PME demonstrated the second highest overall activity when compared to the other tested
enzymes, with specific activities ranging from 6 units (mg protein)-1 to 34 units (mg protein)1
(Fig. 12A). PME activity [(mg protein)-1 basis, Fig. 12A] decreased throughout floral bud
development (Phase I) and flower opening (Phase II), reaching a minimum level of
approximately 9 units (mg protein)-1. There was an approximately three fold increase in
activity as the flowers senesced (Phase III).
61
Phase
I
II
III
Fig. 12. Changes in pectin methylesterase activity in sepals and petals during
development of D. crumenatum floral buds. (A) Activity expressed on (mg
protein)-1 basis, (B) activity expressed on (floral part)-1 basis, (C) activity
expressed on gFW-1 basis. Phases of development – I: floral bud development, II:
flower opening, III: flower senescence. Each value is mean ± SE (n = 5) and
values with different alphabet(s) (sepals: a – d; petals: a’ – d’) are significantly
different according to Fisher’s LSD test (P < 0.05).
62
On a per floral part basis (Fig. 12B), an increase in PME activity was observed during the
early stages of floral bud development (Phase I); it decreased throughout flower opening
(Phase II) and flower senescence (Phase III).
On a per gram fresh weight basis (Fig. 12C), PME activity showed a decreasing trend
during floral bud development (Phase I), and it remained at low and constant levels through
flower opening (Phase II) and flower senescence (Phase III).
4.4.5 β-galactosidase
β-gal exhibited the third highest overall activity when compared to the other tested enzymes,
with specific activities ranging from 2 units (mg protein)-1 to 12 units (mg protein)-1 (Fig.
13A). β-gal activity [(mg protein)-1 basis, Fig. 13A)] remained unchanged and was
approximately 4 units (mg protein)-1 during floral bud development (Phase I) and flower
opening (Phase II). Upon flower senescence (Phase III), a significant increase (P < 0.05) in
β-gal activity, from approximately 4 units (mg protein)-1 to 10 units (mg protein)-1 was
observed.
On a per floral part basis (Fig. 13B), an increase in β-gal activity was observed during
early stages of floral bud development (Phase I); it decreased as development progressed, and
the level of activity increased again during flower senescence (Phase III).
On a per gram fresh weight basis, a drastic decrease in β-gal activity was observed
during floral bud development (Phase I) and flower opening (Phase II), and it subsequently
increased during flower senescence (Phase III) (Fig. 13C).
63
Phase
I
II
III
Fig. 13. Changes in β-galactosidase activity in sepals and petals during development
of D. crumenatum floral buds. (A) Activity expressed on (mg protein)-1 basis, (B)
activity expressed on (floral part)-1 basis, (C) activity expressed on gFW-1 basis.
Phases of development – I: floral bud development, II: flower opening, III: flower
senescence. Each value is mean ± SE (n = 5) and values with different alphabet(s)
(sepals: a – c; petals: a’ – c’) are significantly different according to Fisher’s LSD
test (P < 0.05).
64
4.4.6 β-glucosidase
β-glu exhibited the highest overall activity when compared to the other tested enzymes, with
specific activities ranging from 9 units (mg protein)-1 to 43 units (mg protein)-1 (Fig. 14A).
During late stages of floral bud development (Phase I), β-glu activity [(mg protein)-1 basis,
Fig. 14A] increased sharply by approximately two folds, and decreased during flower
opening (Phase II) and flower senescence (Phase III).
On a per floral part basis (Fig. 14B), an increase in β-glu activity was observed during
floral bud development (Phase I); it subsequently decreased upon flower opening (Phase II)
and flower senescence (Phase III).
On a per gram fresh weight basis (Fig. 14C), a constant decrease in β-glu activity was
observed throughout floral bud development (Phase I), flower opening (Phase II) and flower
senescence (Phase III).
4.4.7 β-mannosidase
Activity of β-man was very much lower compared to that of PME, β-gal and β-glu, with
specific activities ranging from 0.4 units (mg protein)-1 to 1.8 units (mg protein)-1 (Fig. 15A).
β-man activity [(mg protein)-1 basis, Fig. 15A] demonstrated a gradual increase throughout
floral bud development (Phase I), flower opening (Phase II) and flower senescence (Phase
III), with a maximum activity of approximately 1.8 units (mg protein)-1 in the senesced
flower.
On a per floral part basis (Fig. 15B), an increase in β-man activity was observed
during early stages of floral bud development (Phase I), which then decreased over flower
opening (Phase II) and flower senescence (Phase III).
.
65
Phase
I
II
III
Fig. 14. Changes in β-glucosidase activity in sepals and petals during development of
D. crumenatum floral buds. (A) Activity expressed on (mg protein)-1 basis, (B) activity
expressed on (floral part)-1 basis, (C) activity expressed on gFW-1 basis. Phases of
development – I: floral bud development, II: flower opening, III: flower senescence.
Each value is mean ± SE (n = 5) and values with different alphabet(s) (sepals: a – c;
petals: a’ – c’) are significantly different according to Fisher’s LSD test (P < 0.05).
66
Phase
I
II
III
Fig. 15. Changes in β-mannosidase activity in sepals and petals during
development of D. crumenatum floral buds. (A) Activity expressed in mg protein-1
basis, (B) activity expressed in (floral part)-1 basis, (C) activity expressed in gFW-1
basis. Phases of development – I: floral bud development, II: flower opening, III:
flower senescence. Each value is mean ± SE (n = 5) and values with different
alphabet(s) (sepals: a – c; petals: a’ – c’) are significantly different according to
Fisher’s LSD test (P < 0.05).
67
On a per gram fresh weight basis (Fig. 15C), β-man decreased sharply during the late
stages of floral bud development (Phase I), and it increased slightly upon flower senescence
(Phase III).
4.4.8 β-xylosidase
Similar to β-man, activity of β-xyl was very much lower compared to that of PME, β-gal and
β-glu, with specific activities ranging from 0.3 units (mg protein)-1 to 1.9 units (mg protein)-1
(Fig. 16A). β-xyl activity [(mg protein)-1 basis, Fig. 16A] was at low and relatively constant
levels throughout floral bud development (Phase I) and flower opening (Phase II). Upon
flower senescence (Phase III), the activity of β-xyl increased by approximately three to four
folds.
On a per floral part basis (Fig 16B), an increase in β-xyl activity was observed during
early stages of floral bud development (Phase I); it remained constant till flower opening
(Phase II), then decreased upon flower senescence (Phase III).
On a per gram fresh weight basis (Fig. 16C), a constant decrease in β-xyl activity was
observed during floral bud development (Phase I), it remained at low and constant levels
during flower opening (Phase II) and flower senescence (Phase III).
4.5 Expression of cell wall-based enzyme gene transcripts
Based on the results on the activities of the various cell wall-based enzymes investigated, βgal and PME were selected for further studies on the expression of their gene transcripts at
various stages of floral bud/ flower development. Both β-gal and PME have been suggested
to be key enzymes involved in various cell wall modification processes during fruit ripening
and flower opening (Brummell and Harpster 2001; O’Donoghue 2006). Since the results
68
Phase
I
II
III
Fig. 16. Changes in β-xylosidase activity in sepals and petals during development of
D. crumenatum floral buds. (A) Activity expressed in mg protein-1 basis, (B) activity
expressed in (floral part)-1 basis, (C) activity expressed in gFW-1 basis. Phases of
development – I: floral bud development, II: flower opening, III: flower senescence.
Each value is mean ± SE (n = 5) and values with different alphabet(s) (sepals: a – c;
petals: a’ – c’) are significantly different according to Fisher’s LSD test (P < 0.05).
69
from the enzyme activities demonstrated that both sepals and petals exhibited the same
patterns of change (Fig. 12, 13), expression of the gene transcripts were analysed in samples
consisting of sepals and petals together.
4.5.1 Total RNA integrity and quality
Total RNA isolated from D. crumenatum samples from various floral bud/ flower
developmental stages were assessed to be of high integrity and high quality. The RNA
samples from each stage of development exhibited two distinct bands (28S ribosomal RNA
and 18S ribosomal RNA) when examined on agarose gels, indicating that the RNA had high
integrity with minimal degradation (Fig. 17). The A260/280 ratios of the RNA samples were
between 1.63 – 2.02, and A260/230 ratios were above 2.0, indicating the absence of protein
and carbohydrate contamination respectively (Table 5).
4.5.2 Optimization of PCR
Gradient PCR for the control gene β-TUB (Fig. 18A) demonstrated that the optimum
annealing temperature for the pair of primers used was in the range of 50°C – 55°C, and the
primers exhbited high specificity, with only 1 distinct band of size 830 bp. A wide range of
temperatures were suitable as optimum annealing temperature for the pair of primers for βGAL (Fig. 18B), ranging from 50°C – 59°C. The pair of primers was also specific, with only
1 distinct band size of 441 bp. Gradient PCR for PME (Fig. 18C) demonstrated that the
optimum annealing temperature for the pair of primers used was in the range of 48°C – 54°C.
Between 48°C – 50°C, however, a high degree of smearing was obseerved. The PME primers
demonstrated high specificity, with only 1 distinct band of size 292 bp. Since the optimum
annealing temperature for all three pairs of primers coincided within the same temperature
range, 52°C was selected as the final temperature to be used for all further PCR
70
Fig. 17. Electrophoresis of total RNA from sepals and petals of D. crumenatum,
with 28S and 18S ribosomal RNAs indicated.
71
Table 5. Quality of RNA obtained from D. crumenatum samples at various floral bud/
flower developmental stages (n = 4).
Sample
(days after induction)
Ratio of absorbance
260nm/ 280 nm
Ratio of absorbance
260nm/ 230nm
4
1.76 ± 0.16
2.19 ± 0.15
7
1.82 ± 0.15
2.27 ± 0.13
9
1.63 ± 0.09
2.06 ± 0.19
10
1.96 ± 0.19
2.21 ± 0.11
12
2.02 ± 0.12
2.40 ± 0.17
72
°C
°C
°C
Fig. 18. Gradient PCR amplification of (A) β-TUB, (B) β-GAL and (C) PME
transcripts, conducted at various annealing temperatures.
73
amplifications. Amplification of all three genes at various numbers of amplification cycles
showed that their expression levels continued to increase up till at least 32 cycles (Fig. 19).
Further PCR amplifications were thus conducted for 30 cycles for β-TUB, β-GAL and PME.
4.5.3 Expression of β-GAL and PME during floral bud and flower development
β-GAL transcripts were expressed at low levels during floral bud development (Phase I),
which then became highly expressed upon flower opening (Phase II) and flower senescence
(Phase III) (Fig. 20A).
No expression of PME transcripts was observed during the early stages of floral bud
development (Phase I), but PME transcripts were expressed at very low levels during the
later stages of floral bud development. Expression of PME transcripts continued to increase
upon flower opening (Phase II) and flower senescence (Phase III) (Fig. 20B).
4.6 Membrane stability
Significant increases in percentage of electrolyte leakage (P < 0.05) were observed during
early stages of bud development (day 4 – day 9) (Fig. 21A) in sepals and petals, and the
percentage of leakage continued to increase significantly up till senescence stage in the
petals. Membrane stability (determined as membrane stability index, MSI) of sepals and
petals declined with development of D. crumenatum floral buds (Fig. 21B). The highest
decline in membrane stability was observed during early stages of development; it decreased
from approximately MSI 80 on day 4 to MSI 69 on day 7 for sepals, and from MSI 72 on day
4 to MSI 62 on day 7 for petals.
74
Fig. 19. PCR amplifications of β-TUB, β-GAL and PME transcripts
conducted at an annealing temperature of 52°C, and at 24, 26, 28, 30
and 32 cycles each.
75
Phase
Fig. 20. Expression of (A) β-GAL and (B) PME transcripts at various developmental
stages of D. crumenatum floral buds/ flowers. β-TUB was used as the control gene.
Phases of development – I: floral bud development, II: flower opening, III: flower
senescence.
76
Phase
I
II
III
Fig. 21. Membrane stability of sepals and petals of D. crumenatum at various floral
bud/ flower developmental stages. (A) Electrolyte leakage of sepals and petals, (B)
membrane stability index of sepals and petals. Phases of development – I: floral bud
development, II: flower opening, III: flower senescence. Each value is mean ± SE (n
= 5) and values with different alphabet(s) (sepals: a – c; petals: a’ – e’) are
significantly different according to Fisher’s LSD test (P < 0.05).
77
4.7 Control of flower opening
One day after treatment, treated day 9 floral buds exhibited either full flower opening (Fig.
22A), or remained as dormant floral buds with day 9 features showing no signs of senescence
(Fig. 22B), or displayed incomplete flower opening (Fig. 22C, D). Treatments with water,
GA, AOA and AOA + glucose resulted in almost 100 % full flower opening in D.
crumenatum floral buds, while the presence of BA in treatment solutions greatly reduced the
percentage of full flower opening (Table 6). The greatest reduction, of approximately 50 %,
in full flower opening occurred when day 9 floral buds were treated with 10-8 M BA. The
inhibitory effects of BA on full flower opening were also observed in the combined
treatments consisting of both BA and GA. While GA alone resulted in almost 100 % full
flower opening at all the tested GA concentrations, the inclusion of BA resulted in about 15 –
40 % reduction in full flower opening, depending on the concentration of BA used.
The percentages of treated floral buds that remained dormant with day 9 features, and
without any signs of senescence (Fig. 21B), were as shown in Table 7. Dormancy of floral
buds occurred only in the presence of BA, with the highest percentage of approximately 45
% when treated with 10-8 M BA. All of these dormant floral buds aborted without showing
any signs of flower opening. All of the dormant floral buds began to show signs of
senescence, turning transparent and exhibiting signs of discolouration in the pedicel by day
11 (Fig. 22I). The presence of BA also resulted in incomplete flower opening in some floral
buds (Fig. 22C, D). These floral buds did not attain full flower opening and senesced at the
same time and rate as those that displayed full flower opening.
All the fully opened and incompletely opened flowers began to show signs of
senescence on day 11. Except for those treated with AOA + glucose, all flowers senesced
78
similar to the control, with discolouration of the pedicel and the closing or wilting of all
sepals and petals (Fig. 22G). Approximately 25 % of the flowers from the AOA + glucose
treatment exhibited abnormal senescing phenotype on day 11. They remained fully open with
the sepals and petals turning translucent, and were thus still considered as senesced flowers
(Fig. 22E). By day 12, the sepals and petals from these flowers became even more
translucent and turned ‘outwards’, instead of the usual browning and closing of the flowers
(Fig. 22F, H).
79
Fig. 22. General physical features of day 9 D. crumenatum floral buds after
treatments. (A) Fully opened flower, (B) dormant floral bud with day 9 features and
no signs of senescence, (C) incompletely opened flower with slight opening on the
sides and tip of the floral bud, (D) incompletely opened flower, (E) fully opened
flower with translucent sepals and petals, (F) senescent flower turned ‘outwards’
with translucent sepals and petals, (G) wilted flower, (H) magnified view of flower
in F, (I) senescent floral bud. Scale bar = 0.5 cm.
80
Table 6. Percentages of D. crumenatum floral buds that displayed full flower opening one day
after treatment. Treatments were applied on day 9 floral buds that normally proceeded to
flower opening the next day.
Treatment
Total number of
floral buds
20
Number of fully
opened flowers
20
Percentage
flowered (%) ± SE
100 ± 0
BA (10-6 M)
20
11
55 ± 22
BA (10-8 M)
20
10
50 ± 6
BA (10-10 M)
20
12
60 ± 14
GA (10-6 M)
20
19
95 ± 5
GA (10-8 M)
20
20
100 ± 0
GA (10-10 M)
20
19
95 ± 5
BA/GA (10-6 M)
20
13
65 ± 10
BA/GA (10-8 M)
20
12
60 ± 8
BA/GA (10-10 M)
20
17
85 ± 9
AOA (10-6 M)
20
20
100 ± 0
AOA (10-8 M)
20
20
100 ± 0
AOA (10-6 M) + 0.1 M
glucose
20
20
100 ± 0
AOA (10-8 M) + 0.1 M
glucose
20
20
100 ± 0
Distilled water (control)
81
Table 7. Percentages of D. crumenatum floral buds that displayed dormancy after one day of
treatment, and percentages of dormant floral buds that subsequently aborted two days after
treatment. Floral buds were considered to be dormant if they maintained the typical features of
a normal day 9 floral bud as described in Table 1, without any signs of senescence.
Treatment
Distilled water (control)
Total
number of
floral buds
20
Number of
white floral
buds
0
Percentage of
dormant floral
buds (%) ± SE
0
Percentage of
aborted floral
buds (%)
0
BA (10-6 M)
20
3
15 ± 5
100
BA (10-8 M)
20
9
45 ± 6
100
BA (10-10 M)
20
2
10 ± 6
100
GA (10-6 M)
20
0
0
0
GA (10-8 M)
20
0
0
0
GA (10-10 M)
20
0
0
0
BA/GA (10-6 M)
20
4
20 ± 11
100
BA/GA (10-8 M)
20
2
10 ± 6
100
BA/GA (10-10 M)
20
0
0
0
AOA (10-6 M)
20
0
0
0
AOA (10-8 M)
20
0
0
0
AOA (10-6 M) + 0.1 M
glucose
20
0
0
0
AOA (10-8 M) + 0.1 M
glucose
20
0
0
0
82
Chapter 5. Discussion
The cell wall is a dynamic structure that can undergo diverse changes during different
developmental phases of the plant. Many important physiological functions are associated
with cell wall modifications or cell wall remodelling. These include structural and
mechanical support, maintenance of cell shape, control rate of growth, protection against
dehydration, defence against pathogens and cell-cell interactions (Carpita and Gibeaut 1993;
Cosgrove 1999; Carpita and McCann 2000). While the roles of cell wall modifications have
been studied in great detail in various plant processes such as seed germination (Edwards et
al. 1985; Crombie et al. 1998; Buckeridge et al. 2000; Tine et al. 2000), abscission
(Lashbrook et al. 1994; Patterson 2001; Fulton and Cobbett 2003) and fruit ripening
(Brummell and Harpster 2001; Brummell 2006), the association of cell wall remodelling and
flowering remains rather unclear, with limited studies conducted to date. The main interest
was in determining if there were distinctive changes, occurring in sepals and petals of D.
crumenatum floral buds and flowers, which might contribute to the opening and senescing of
the flowers.
5.1 Cell wall changes related to D. crumenatum floral bud/ flower development
The development of D. crumenatum floral buds and opening of the flowers were found to be
accompanied by increasing disorganization of parenchyma cells and appearance of
intercellular spaces (Fig. 5). It was hypothesized that such anatomical changes were due to
alterations in cell wall constituents that commenced in young floral buds, and that such cell
wall modifications might regulate the development, opening and senescing of floral
buds/flowers.
83
The ‘looseness’ observed in the structures of sepals and petals during development of
D. crumenatum floral buds (Phase I) (Fig. 5) appeared to be due to the breakdown of cell
wall materials. On the contrary, the results obtained demonstrated that the net amount of cell
wall materials (in the form of ethanol-insoluble residue, EIR) increased during floral bud
development, which was proportional to the increase in weight and size of the sepals and
petals. Loss of cell wall materials occurred only during flower opening (Phase II) and flower
senescence (Phase III) (Fig. 4, 7A, 8A). The synthesis of cellulose, the principal scaffold in
cell walls, was observed to increase in conjunction with the increase in cell wall materials
during floral bud development (Fig. 4, 7B, 8B). The net amount of cellulose subsequently
decreased slightly during flower senescence, indicating cellulose hydrolysis. However,
enzymatic analyses demonstrated the absence of cellulase activity at all developmental stages
(Fig. 11). It was possible that cellulase was present in sepals and petals of D. crumenatum,
but its activity was at very low and undetectable levels, or carboxymethylcellulose and filter
paper could be unsuitable substrates for cellulase in D. crumenatum. Substrate specificities of
cellulases have been studied with proteins purified from avocado, strawberry and tomato, and
results demonstrated that different substrates resulted in varying levels of cellulase activity
within each species (Hayashi et al. 2005; Urbanowicz et al. 2007).
The net quantity of hemicelluloses increased during D. crumenatum floral bud
development and decreased during flower opening and flower senescence (Fig. 4, 7C, 8C).
The results indicated that although hemicelluloses were synthesized during floral bud
development, the re-distribution of hemicelluloses due to increase in size of sepals and petals
was not in synchrony with the rate of synthesis. As a result, there existed a dilution of
hemicelluloses and a decrease in concentration when expressed on a per floral part basis (Fig.
84
7C). Hemicelluloses play important structural roles in the cell walls by contributing to the
cross-linking of cellulose microfibrils, creating a network between the various microfibrils
(Carpita and McCann 2000). One of the most significant networks, the cellulose-xyloglucan
network, has been suggested to be a major load-bearing structure in the cell walls, and is an
important constraint to cell wall loosening (Rose and Bennett 1999). The redistribution and
breakdown of hemicelluloses may result in disassembling or relaxation of the cellulosexyloglucan network, causing cell wall swelling and altering the movement of cell wall
enzymes within the cell wall matrix, varying the access to their respective substrates (Rose
and Bennett 1999; Brummell 2006).
The breakdown of hemicelluloses can be attributed to the actions of various
glycosidases (de Vetten et al. 1991; Minic and Jouanin 2006). The specific activities of β-gal
(Fig. 14A), β-man (Fig. 15A) and β-xyl (Fig. 16A), together with the changes in amount of
soluble proteins with development of D. crumenatum floral buds and flowers (Fig. 9),
resulted in the upregulation of the enzyme activities [units (floral part)-1] during the later
stages of floral bud development (Fig. 14B, 15B, 16B), and might account for the subsequent
breakdown of hemicelluloses during flower opening and senescence. The family of
glycosidases is known to play a crucial role in the degradation of various cell wall
polysaccharides, allowing the remodelling of the cell wall structure (Minic 2008). The
breakdown of cell walls, catalysed by glycosidases, for the mobilization of cell wall storage
polysaccharides in germinating or post-germinating seeds have been widely studied
(Buckeridge et al. 2000). In guar (Cyamopsis tetragonolobus), fenugreek (Trigonella
foenum-greacum) and carob (Ceretonia siliqua), the degradative effects of β-man has been
shown to be important and associated with galactomannan mobilization, following
85
germination of the seeds (Reid 1971, 1985; McClendon et al. 1976). The mobilisation of
xyloglucan in seeds of Tropaeolum majus and Copaifera langsdorffi has also been shown to
be accompanied by the rise in β-gal activity (Edwards et al. 1985; Buckeridge et al. 1992).
Softening of fruits during ripening has been associated with disassembly of the cell wall due
to changes in hemicellulose structure or content. In the ripening of avocado, Japanese pear
(Pyrus pyrifolia) and strawberry (Ronen et al. 1991; Itai et al. 1999; Martinez et al. 2004),
increases in β-xyl activities have been shown to be related to the degradation of arabinan and
xylan during the softening processes.
Pectin hydrolysis in the development of D. crumenatum floral buds and flowers were
not observed (Fig. 4, 7D, 8D). In support of this observation, activities of the enzyme that
catalyses pectin hydrolysis, PG, was also undetected throughout development of the orchid
floral buds/flowers. Pectins are heteropolysaccharides in the cell wall, and are important in
regulating wall porosity, cell-cell adhesion at the middle lamella and movement of enzymes
within the cell wall matrix (Baron-Epel et al. 1988; Pena and Carpita 2004; Brummell 2006).
Degradation of cell wall pectins in the context of fruit ripening and softening has been widely
studied. In the ripening of carambola and grapes, decreases in pectin content was
accompanied by increases in PG activity (Chin et al. 1999; Deng et al. 2005). Pectin
hydrolysis is, however, not the only pectin modification that can result in the remodelling of
the cell wall structure. Pectin solubilisation, resulting from changes in the types of bonds
and/or bond strengths existing between pectin molecules and/ or other matrix molecules, has
been correlated with cell wall swelling, increasing the accessibility of cell wall enzymes to
their respective substrates (Redgewell et al. 1997; Brummell 2006). Water-soluble and
CDTA-soluble pectins are relatively weakly bound to cell wall polysaccharides by molecular
86
entanglements, hydrophobic forces, weak ionic bonds or ionic calcium bridges, while
Na2CO3-soluble pectins are more strongly attached to the cell wall via covalent bonds
(Brummell 2006). Increases in pectin solubilisation have been demonstrated in the ripening
of tomato and avocado that were accompanied by changes in cell wall structure (Carrington
et al. 1993; Wakabayashi et al. 2000). In D. crumenatum, pectin solubilisation changed as
the floral buds and flowers developed. There were increasing proportions of pectins that were
susceptible to the solubilising agents during floral bud development (Phase I), and decreased
pectin solubility during flower opening (Phase II) and senescence (Phase III) (Table 3, 4).
PME, which de-esterifies pectins, has been suggested to increase pectin solubilisation
by creating electronic repulsion between negatively-charged molecules that could result in
the loosening of weakly attached pectins from the cell wall (Grignon and Sentenac 1991). In
D. crumenatum, the specific activity of PME (Fig. 12A) together with changes in the quantity
of soluble proteins (Fig. 9), resulted in a significant increase in PME activity [units (floral
part)-1] during the early stages of floral bud development (Phase I) (Fig. 12B). This
maximum level of PME activity coincided with the largest increase in pectin solubility
(Table 3, 4). However, PME transcript expression could not be related to the activity profile
(Fig. 20B). PME might be coded by more than one gene in D. crumenatum such that the
enzyme activity profile obtained was a quantification of a composite of PME proteins from
two or more highly homologous genes. Studies have demonstrated that PME isoforms in cell
walls are encoded by a multigene family, and the analysis of the genome sequence in
arabidopsis has identified 67 PME-related genes PME (The Arabidopsis Genome Initiative
2000; Micheli 2001). In tomato, PME has also been shown to consist of at least four genes,
some of which are highly homologous (Harriman et al. 1991; Hall et al. 1994; Turner et al.
87
1996; Gaffe et al. 1997). Cell wall modifications, occurring during the ripening of carambola
and grapes, were the result of an increase in PME activity, which was in synchrony with
pectin solubilisation (Chin et al. 1999; Deng et al. 2005). The activity of PME resulting in
increased pectin solubilisation has also been studied during cell separation of root border
cells in pea (Peasum sativum) (Stephenson and Hawes 1994). It was suggested that pectin
solubilisation in the middle lamella caused changes in cellular adhesion and resulted in
separation of the cells. Furthermore, increase in PME activity has been shown to accompany
dormancy breakage and germination of yellow cedar seeds (Chamaecyparis nootkatensis),
and although the precise role of the enzyme in regulating seed germination remains
unknown, the actions of PME must had altered and weakened the cell walls of the
megagametophyte, aiding radicle protrusion and seed germination (Ren and Kermode 2000).
There are many other factors that may also regulate pectin solubilisation and other
polymer modifications. One such candidate could be β-gal, the enzyme responsible for the
degradation of cell wall galactan, regulating cell wall flexibility, intercellular connections
and cell wall porosity, and affecting the mobility of enzymes within the cell wall matrix
(Brummell and Harpster 2001; Brummell 2006). In vitro treatments of cell wall preparations
from papaya with β-gal resulted in increased pectin solubilisation (Ali et al. 1998). In D.
crumenatum, specific activity of β-gal (Fig. 13A) together with changes in the quantity of
soluble proteins (Fig. 9), resulted in significant increases in β-gal activity [units (floral part)1
] during the early stages of floral bud development (Phase I) and flower senescence (Phase
III) (Fig. 13B). β-gal transcript expression, however, peaked only during flower opening
(Phase II) and flower senescence (Phase III) (Fig. 20A). Similar to PME, β-gal might also be
encoded by more than one homologous gene in D. crumenatum. In tomato, at least seven β-
88
gal genes were expressed during development and fruit ripening (Smith and Gross 2000).
Post-transcriptional and post-translational modifications may also result in the differences
between the enzyme activity profile and the expression of the gene transcript.
It is possible that non-enzymic mechanisms might also be involved in cell wall
modifications. In tomato, ascorbate-generated hydroxyl radicals demonstrated non-enzymic
scission of cell wall polysaccharides, and caused an increase in pectin solubilisation
(Dumville and Fry 2003). In the study, physiological concentrations of ascorbate gradually
solubilised pectins present in the cell wall materials of the fruit. In the presence of DMSO,
which is a scavenger for hydroxyl radicals, pectin solubilisation was inhibited, demonstrating
that the mechanism of action of ascorbate was via hydroxyl radicals (Dumville and Fry
2003).
5.2 Model for cell changes accompanying D. crumenatum floral bud/ flower
development
The data obtained suggested that cell wall modifications determining floral bud development,
flower opening and flower senescence in D. crumenatum were accompanied by
modifications in both the cellulose-hemicellulose networks and pectin networks. The results
suggested that temporal changes in cell wall modifications occurred in two stages (Fig. 23).
An early stage corresponding to floral bud development (Phase I), that is characterised by
cellulose synthesis, slight hemicellulose synthesis and major pectin solubilisation. Although
net hemicellulose content increased during this period, the re-distribution of the
polysaccharides due to increases in size of sepals/ petals resulted in an overall dilution of
hemicellulose content per floral part (Fig. 7C), and could have resulted in the loosening of
89
Fig. 23. Proposed model for cell wall modifications accompanying D. crumenatum
floral bud and flower development. Phases of development – I: floral bud development,
II: flower opening, III: flower senescence. Black bars indicate the occurrences of the
particular events.
90
the cellulose/ hemicellulose networks. It was also possible that pectin solubilisation, which
occurred at significant levels during this developmental phase could have loosen the pectin
networks, enhancing enzyme mobilities within the cell wall matrix, resulting in subsequent
cell wall changes. Pectins are the main components of the middle lamella (Brummell 2006;
Liepman et al. 2007). Thus, the occurrence of high pectin solubilisation could also indicate
the dissolution of the middle lamella, resulting in decreased cell adhesion. The second stage
of cell wall modifications coincided with flower opening (Phase II) and flower senescence
(Phase III) (Fig. 23), with extensive breakdown of cellulose and hemicellulose. These
degradative effects probably resulted in the almost total dissolution of the cell walls and
collapse of the parenchyma layers upon senescence of the D. crumenatum flowers (Fig. 5E).
5.3 Species-specific variations in cell wall modifications associated with flowering
The influence of cell wall modifications in the regulation of flowering has only been studied
in a few species of flowers, including carnation, sandersonia and daylily. Cell wall
modifications in petals of carnation and sandersonia were studied in great detail, with known
cell wall composition and cell wall enzyme activities, whereas there were no published data
on cell wall composition of daylily (de Vetten and Huber 1990; de Vetten et al. 1991;
Panavas et al. 1998; O’Donoghue et al. 2002, 2005).
The different species of flowers differ in their time-frames of flower development, i.e.
rate of floral bud development, time of flower opening, duration of flower opening processes,
longevity of flowers (Fig. 24). Of the tested species, carnation has the longest period of floral
bud development, flower opening process and longevity, while daylily has the shortest
91
Fig. 24. Comparison of flower development events in carnation (de Vetten and Huber 1990),
sandersonia (Eason and Webster 1995; O’Donoghue et al. 2002), daylily (Biesleki and Reid
1992; Panavas et al. 1998) and D. crumenatum. Time of fully opened flowers was denoted as
day 0 for all species.
92
developmental period. Due to the variations in the time-frames of flower development of
these flowers, comparisons of flower regulatory factors at each pre-defined developmental
phase (floral bud development – Phase I; flower opening – Phase II; flower senescence –
Phase III) are possible, but not for comparisons at each time point of development.
Comparisons of the cell wall changes between the flowers of different species show
that they share some features, but differ in others (Table 8). In general, cell wall
modifications in the forms of alterations in cell wall composition and cell wall enzyme
activities occurred during development of all the flowers. Increases in cellulose deposition
during flower opening (Phase II) and increases in β-gal activities (presumably resulting in
loss of galactose and changes in pectin networks) during flower senescence (Phase III) have
been observed in all four species. One of the most significant differences between the flowers
was the presence of cellulase and PG in daylily, while both enzymes were absent (or at very
low and undetectable levels) in all the other three species.
There appears to be a species-specific variation in the pattern of cell wall
modifications during flower development, but a complete comparison is difficult owing to
the lack of cell wall composition data from daylily and also the lack of data on various other
cell wall modifications (e.g. activities of β-glu, β-man and β-xyl) from other species. Speciesspecific variation in cell wall modifications has also been reported in ripening fruits. Pectin
solubilisation in ripening avocado was observed to be three times more than in kiwifruit
(Actinidia deliciosa), five times more than in blackberry (Rubus fruticosus), nine times more
than in persimmon (Diospyros kaki) and plum (Prunus domestica), and 12 times more than in
tomato and strawberry; whereas pectin solubilisation was absent or occurred at very low and
insignificant levels in apple (Malus domestica cv. Cox’s Orange Pippin and Malus domestica
93
Table 8. Summary of modifications of cell wall polysaccharides (per floral part basis) and cell wall
enzyme activities (per floral part basis or per mg protein basis) during floral bud development (Phase
I), flower opening (Phase II) and flower senescence (Phase III) of D. crumenatum, carnation (de
Vetten and Huber 1990; de Vetten et al. 1991), sandersonia (O’Donoghue et al. 2002) and daylily
(Panavas et al. 1998). Analysis of cell wall composition and enzymatic activities during Phase I were
not conducted in carnation and sandersonia. Analysis of cell wall composition was not conducted in
daylily throughout development. Symbols*, abbreviations† and legend# are outlined in footnote.
Phase I
(Young floral bud –
Mature floral bud)
Phase II
(Mature floral bud –
Mature flower)
Phase III
(Mature flower – Senesced
flower)
Composition
Cellulose
Hemicellulose
Total pectin
WSP
CSP
SSP
+
+
+
+
+
+
Ø
+ +
+ X
- +
Ø
- +
- X
- +
-
X
-
X
Ø
Ø
+ Ø
+ Ø
Ø
Ø
+ - -
X
X
Enzyme
activity
Cellulase
PG
PME
̀β-gal
β-glu
β-man
β-xyl
Ο
Ο
Ο
Ο
Ο Ο X Ο +
Ο Ο Ο Ο Ø
Ο Ο X Ο
Ο Ο Ο Ο
+- +- Ø
+ +
+- +
+Ø Ø
Ø - X - Ø Ø X Ø Ø
+
-
- - +
Ø +
X X
X
X
X
X
X
X
X
+
+
+
+
+
X + + + +
+ X X
+ X X
X
X
X
*Symbols – +: factor increased in quantity or activity during the developmental phase; -: factor
decreased in quantity or activity during the developmental phase; Ø: factor assayed and remained
unchanged; Ο: factor assayed and absent; X: factor not assayed.
†
Abbreviations – WSP: water-soluble pectin; CSP: CDTA-soluble pectin; SSP: Na2CO3-soluble
pectin.
#
Legend – various font colours indicate different flower species and their respective expression units:
D. crumenatum (per floral part basis); D. crumenatum (per mg protein basis); carnation (per
floral part basis); sandersonia (per floral part basis); daylily (per mg protein basis).
94
cv. Braeburn), watermelon (Citrullus lanatus cv. Charisma) and Nashi pear (Pyrus serotina
cv. Nijisseiki) (Redgwell et al. 1997). The species-specific variations in pectin solubilisation
in these fruits were correlated to cell wall swelling and were suggested to result in the
differences in the texture of the fruits (Redgwell et al. 1997).
5.4 Relationship between flowering and senescence
Flower senescence, which is the end point of flower life, is often a rapid and
synchronous process, and has frequently been linked to the various catabolic and ‘death’
processes that occur after flower opening (O’Donoghue 2006). Visible markers of flower
senescence often include wilting, sepal/ petal abscission, browning and drying (O’Donoghue
2006). Physiological changes associated with flower senescence include membrane
degradation, alterations in cell wall structures, protein remobilization and DNA laddering
(Rubinstein 2000; Wagstaff et al. 2003; Zhou et al. 2005).
Previously, studies on carnation and sandersonia suggested that just prior to flower
opening (late floral bud stage), a certain degree of cell wall dismantling had already started
(de Vetten and Huber 1990; de Vetten et al. 1991; O’Donoghue et al. 2002).
In D.
crumenatum, senescence-related events such as cell wall modifications occurred as early as
during young floral bud development (Fig. 7, 8, Table 8). Also, membrane stability began to
decrease during the period of floral bud development (Fig. 21), indicating the breakdown of
the cell membrane. The data suggested that the onset of flower senescence occurred as early
as during floral bud development. Early commencement of flower senescence was also
observed in morning glory (Ipomoea tricolor), where alterations to cell shape and cell wall
thickness occurred even before flower opening (Phillip and Kende 1980). Petal senescence
95
was also shown to begin extremely early in Alstromeria peruviensis var. Samora (Wagstaff et
al. 2003). In the study, indicators of senescence such as DNA laddering and nuclear
degradation appeared as early as two days before the onset of flower opening, and these
senescence processes proceeded throughout flower opening and the eventual visible
senescence of the flowers (Wagstaff et al. 2003). Pollination and fertilization of flowers
promote sepal and petal senescence, while keeping the fertilized ovaries viable; in nonpollinated and unfertilized flowers, whole flowers senesce and die (van Doorn 1997). The
possible onset of senescence, prior to flower opening or in non-pollinated/ unfertilized
flowers, would thus infer a modification of the senescence programme, due to a cascade of
signals generated upon pollination and/ or fertilization that results in the senescence of the
sepals and petals but not the ovaries.
5.5 Effects of growth regulators on the control of flower opening
The roles of PGRs in controlling various plant growth processes has been intensely studied
and many have been identified as putative floral signals (van Doorn and van Meeteren 2003;
Corbesier and Coupland 2006; Sim et al. 2008). These include cytokinins and gibberellin
(Corbesier and Coupland 2006). Cytokinins and gibberellins have also been shown to be
capable, through their interactions, of regulating floral evocation and floral bud/ flower
development (Bernier 1988; Day et al. 1995; Setyadjit et al. 2004; Kim and Miller 2008). It
is interesting that out of all the treatments used in the study, reductions in percentages of
flower opening and signs of floral bud dormancy occurred only in the presence of BA (Fig.
22, Table 6, 7). All of these BA-induced dormant floral buds eventually aborted, with none
proceeding on to flower opening (Table 7). The data suggested that BA (at least at the tested
96
concentrations) had the capacity to induce certain signals cascade(s) that altered the
physiological processes and/or biochemical processes and/or genetic regulatory pathways
that usually result in the normal flowering phenotype of D. crumenatum. On the other hand,
BA (at least at the tested concentrations) was also too potent, causing irreversible changes or
damages to the various regulatory processes and/or pathways, finally resulting in the abortion
of flowering.
Similar to the results obtained from D. crumenatum, exogenous application of BA to
harvested Grevillea Sylvia inflorescences resulted in a decrease in percentages of flower
opening (Setyadjit et al. 2004). Furthermore, in Boronia heterophylla, treatment of the plants
with BA resulted in delayed flowering (Richards 1985). It appears that flower opening is
suppressed by exogenous BA. In Cosmos sulphureus, endogenous cytokinin concentrations,
in particularly zeatin and zeatin riboside, were shown to be at low levels during initial floral
bud development, increased prior to flower opening, and decreased upon full bloom (Saha et
al. 1985). In Boronia megastima, the rapid floral bud development stages were accompanied
by increases in cytokinin concentrations, in particularly zeatin, and decreases in carbohydrate
concentration; upon anthesis, zeatin concentrations decreased while zeatin riboside
concentrations increased (Day et al. 1995). Cytokinins have been suggested to regulate floral
bud/ flower development by controlling the metabolism and distribution of carbohydrates to
the floral buds/ flowers (Munoz et al. 1990; Zieslin and Khayat 1990). The influence of
carbohydrates distribution on cytokinin synthesis may also be possible (Day et al. 1995).
Previous studies have demonstrated that exogenous application of BA to raceme
tissues of soybean (Glycine max L. Merr.) prevented flower abortion (Reese et al. 1995;
Nagel et al. 2001). It was suggested that the action of BA involved redirecting the
97
movements of assimilates and resources to the treated floral buds, thus increasing the sink
strength and growth rates of the floral buds, and consequently reducing abortion of flowering
(Reese et al. 1995; Nagel et al. 2001). In D. crumenatum, however, exogenous BA was
found to have flower abortion-inductive effects (Table 7). To our knowledge, this is the first
report on the ability of exogenous BA to cause abortion of flowering during the final stages
of floral bud development. In the opening of flowers, carbohydrate traffic and osmolarity
status are considered important driving forces behind petal movements (van Doorn and van
Meeteren 2003). It could be possible that the exogenous application of BA to excised D.
crumenatum floral buds, just one day prior to flower opening, caused severe changes to the
partitioning and metabolism of carbohydrates in the floral buds. Such changes would result in
incomplete sepal/ petal expansion that is required to evoke floral bud crack and flower
opening (van Doorn and van Meeteren 2003). Further investigations are required to elucidate
this possibility. The data also suggested that flower opening and anthesis are active processes
that involve the fine concerted balance of various regulators. In D. crumenatum floral buds
that were already committed to anthesis, alterations to this balance as late as during the final
stages of floral bud development, just prior to the onset of flower opening, might have caused
total disruption to the flower opening processes.
While exogenous application of GA to harvested D. crumenatum floral buds
demonstrated no effects on flower opening, treatment of excised iris (Iris x hollandica cv.
Blue Magic) with GA promoted flower opening (Celikel and van Doorn 1995). Similarly,
pulsing with GA in cut Polianthes tuberosa L. cv. Double also enhanced flower opening (Su
et al. 2001). It was suggested that GA-stimulated ethylene production resulted in the
promotion of flower bud opening (Su et al. 2001). In D. crumenatum, the applications of BA
98
or GA, alone or in combinations, produced different results in the reduction of flower
opening and induction of bud dormancy (Table 6, 7). The stimulatory or inhibitory effects
that BA and GA could have on each another were also demonstrated in the flowering of
Miltoniopsis orchid hybrids (Matsumoto Brower 2006). In the study, inhibitory effects of BA
on flowering were rescued by the addition of GA (Matsumoto Brower 2006). Furthermore, in
Paphiopedilum (Macabre x glanduliferum), the stimulatory effects of GA on flowering were
shown to be reduced by BA (Miguel et al. 2008).
AOA + glucose treatments have been shown to promote flower opening and suppress
senescence in D. ‘Jew Yuan Tew’ inflorescences (Rattanawisalanon et al. 2003). Longevity
of pollinated D. ‘Heang Beauty’ flowers also increased with AOA + glucose treatments
(Chandran et al. 2006). In D. crumenatum, however, effects of AOA + glucose on flower
opening and senescence of flowers were not observed (Table 6). The differences in AOA +
glucose effects observed in the previous two studies could be due to the presence of more
sinks (floral buds/ flowers), resulting in partitioning and dilution of AOA and/ or glucose,
and pollination, resulting in changes in overall physiology, respectively. It is interesting that
AOA + glucose treatments resulted in an abnormal senescing phenotype in treated D.
crumenatum floral buds, where the sepals and petals turned translucent but remained fully
open (Fig. 22E, F, H). AOA has been proposed to act as an anti-microbial agent, enabling the
continuous uptake of water in flowers (Rattanawisalanon et al. 2003; Chandran et al. 2006).
The resultant increases in water and glucose contents in D. crumenatum could have resulted
in the maintenance of the fully open stance, even in senescing flowers.
99
5.6 Further works
Cell wall modifications are very diverse (Brummell 2006), and it is highly possible that other
cell wall changes, such as depolymerization of hemicelluloses and pectins, occur and are
important in regulating floral bud development, flower opening and senescence in D.
crumenatum. The presence and activities of various other cell wall enzymes, such as
glycanases and expansin, also remain to be elucidated. The identification of these cell wall
modifications would provide a clearer understanding of the importance and roles that cell
walls have in regulating flower development. Also, investigating the possible roles of nonenzymic mechanisms in controlling cell wall alterations could provide further insight to the
involvement of the cell wall in controlling flowering. Whether variations in patterns of cell
wall modifications are the cause for the diversity in the rates of floral bud development, time
of flower opening and longevity of the flowers in different species remain to be investigated.
Furthermore, determining the presence of changes in patterns of cell wall modifications
between BA-induced dormant/aborted floral buds and untreated wild-type floral buds may
provide useful information on the role of BA in controlling flower development in D.
crumenatum.
100
Chapter 6. Conclusion
Co-ordinated cell wall modifications, such as alterations in specific cell wall components and
activities of cell wall-based enzymes, have important roles in the regulation of floral bud
development, flower opening and flower senescence in D. crumenatum. In particular, floral
bud development in D. crumenatum appeared to be accompanied by the loosening of
cellulose/ hemicellulose and pectin networks, and solubilisation of the middle lamella. Such
events probably resulted in decreases in cell-cell adhesion, loosening of the cell wall matrix,
and increases in mobility of cell wall-based enzymes and their access to the respective
substrates. In D. crumenatum sepals and petals, breakdown or total hydrolysis of the cell wall
only occurred upon flower opening and flower senescence. Expression of the cell wall
enzyme transcripts, however, did not correspond with the activity profiles of the enzymes,
and this might be explained by the presence of more than one gene coding for each enzyme.
Senescence-indicative events such as cell wall modifications and decreases in membrane
stability were found to commence during early floral bud development in D. crumenatum,
suggesting the early onset of a senescence programme.
This study demonstrated that flower opening in mature D. crumenatum floral buds
could be reduced, halted or aborted in the presence of exogenous BA. The importance of the
cytokinin in regulating floral bud development, flower opening, anthesis and the various
processes which precede senescence, and the mechanism through which it alters flower
opening remains to be determined. Specifically, whether BA induced delayed and/ or aborted
flowering occurred due to changes in patterns of cell wall modifications is of interest.
101
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Appendix A: Publication in Scientia Horticulturae
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Contents lists available at ScienceDirect
Scientia Horticulturae
journal homepage: www.elsevier.com/locate/scihorti
Regulation of flower development in Dendrobium crumenatum by changes in
carbohydrate contents, water status and cell wall metabolism
You-Min Yap, Chiang-Shiong Loh, Bee-Lian Ong *
Department of Biological Sciences, National University of Singapore, 14 Science Drive 4, Singapore S117543, Singapore
A R T I C L E I N F O
A B S T R A C T
Article history:
Received 4 March 2008
Received in revised form 5 June 2008
Accepted 23 June 2008
The involvement of carbohydrates, water potential, cell wall components and cell wall-based enzymes in
regulating flower development in Dendrobium crumenatum was investigated. Plants were subjected to
cold treatment to release floral buds from dormancy, and the various parameters were investigated from
young floral bud stage till flower senescence. Development of floral buds was accompanied by
progressive decrease in concentrations of fructans and starch. Upon full flower opening, concentration of
soluble sugars was maximum, accompanied by a more negative water potential. High pectin
methylesterase activity was observed during early bud development and decreased thereafter.
Significant increase in activities of b-galactosidase, b-mannosidase and b-xylosidase was also observed
during floral bud development. The cell walls of sepals and petals were modified extensively during floral
bud and flower development, as observed by changes in the amounts of celluloses, hemicelluloses and
total pectin. Pectin solubilisation was also observed to commence during early floral bud development.
These results indicated that carbohydrate hydrolysis, osmotic changes and cell wall dissolution that
began early in young floral buds, all regulated flower development in this sympodial orchid. Possible
applications of the findings in the horticultural industry are discussed.
ß 2008 Published by Elsevier B.V.
Keywords:
Orchid
Flower development
Carbohydrates
Osmolality
Cell wall composition
Cell wall hydrolases
1. Introduction
Flowering is a critical event in the life-cycle of angiosperms,
allowing for the reproduction of these plants. Flowers of various
plants are also highly prized objects of beauty and are commercially valuable. Hence, understanding the various processes that
regulate flower opening and senescence could help to enhance the
visual quality and vase-life of flowers, and thus increasing their
commercial value. Besides, with the world’s increasing interest in
‘green buildings’ to aid energy efficiency and the accompanying
issue of using flowers for aesthetic benefits (Spala et al., 2008),
understanding flower physiology is very important. However,
publications on the physiology of tropical flowers are limited and
the few detailed studies focus mainly on flowers of temperate
species such as carnation (Dianthus caryophyllus L.), daylily
(Hemerocallis spp.), Asiatic lily (Lilium hybrid), rose (Rosa) and
sandersonia (Sandersonia aurantiaca (Hook.)).
In the opening of flowers, changes in carbohydrate metabolism
and cell osmolarity are considered important driving forces behind
petal movements (van Doorn and van Meeteren, 2003). Rapid
* Corresponding author. Tel.: +65 65162852; fax: +65 67792486.
E-mail address: dbsongbl@nus.edu.sg (B.-L. Ong).
0304-4238/$ – see front matter ß 2008 Published by Elsevier B.V.
doi:10.1016/j.scienta.2008.06.029
flower opening in many species, including roses (Ho and Nichols,
1977), daylily (Bieleski, 1993), Asiatic lilies (Bieleski et al., 2000)
and creeping bellflowers (Campanula rapunculoides) (Vergauwen
et al., 2000), was related to the hydrolysis of reserve carbohydrates.
Rapid petal movements are also highly correlated with cell sap
osmolarity changes, which regulate the direction of water movements, resulting in turgor changes and cell expansion (Ho and
Nichols, 1977; Hew et al., 1989; Bieleski, 1993). Changes in
carbohydrate metabolism and cell sap osmolarity are, therefore,
intimately linked in the process of petal expansion and flower
opening.
In recent years, besides the roles of carbohydrates and cell
osmolarity, there has been some focus on the possible involvement
of cell wall metabolism in the regulation of flowering (de Vetten
and Huber, 1990; O’Donoghue et al., 2002). These studies also
addressed the question on whether flowering could be a process
induced by the senescence programme of the plant and/or specific
plant organ. Some supporting evidences included the upregulation
of a phosphoenolpyruvate mutase mRNA, involved in regulating
hydrolytic enzymes that resulted in membrane degradation in
senescing carnation petals (Wang et al., 1996), the upregulation in
levels of a mRNA that codes for proteins controlling the oxidation
of membrane lipids, prior to or during flower senescence in daylily
(Panavas et al., 1999), and the increasing trend of DNA laddering
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Y.-M. Yap et al. / Scientia Horticulturae 119 (2008) 59–66
throughout petal development in Alstroemeria (Wagstaff et al.,
2003). While molecular evidence is increasing, structural and
physiological evidences are quite limited. Studies on carnation and
sandersonia flowers demonstrated that the transition of floral
stages from opening to fully mature flower till senescence was
accompanied by changes in the levels of various cell wall polymers
such as cellulose and pectins, and activities of cell wall-based
enzymes (de Vetten and Huber, 1990; O’Donoghue et al., 2002).
These observations were similar to the loss of cell wall integrity in
ripening fruits of carambola (Averrhoa carambola) and grapes (Vitis
vinifera) (Chin et al., 1999; Deng et al., 2005). In daylily flowers,
analyses of cell wall composition were not published, but reported
changes in activities of cell wall-based enzymes during flower
development suggested the involvement of cell wall metabolism in
flowering (Panavas et al., 1998). While cellulase activity was
detected in daylily flowers, it was reported to be absent in
sandersonia flowers, indicating the possibility of a species-specific
variation in cell wall metabolism that regulates flowering
(O’Donoghue et al., 2002; Panavas et al., 1998). In the abovementioned studies on carnation and sandersonia flowers, cell wall
changes were compared only between stages of late bud (just prior
to opening), opening flower, mature flower, wilting flower and
senesced flower (de Vetten and Huber, 1990; O’Donoghue et al.,
2002). To fully understand if flowering is a consequence of a
senescence programme that has already started, investigating the
physiological changes occurring throughout the development of a
newly induced young floral bud till flower senescence would be
advantageous. Pollination and fertilisation of flowers promote
sepal/petal senescence, while keeping the fertilised ovary viable;
in non-pollinated and unfertilised flowers, whole flowers senesce
and die (van Doorn, 1997). The possible onset of senescence prior
to flower opening would thus infer a modification of the
senescence programme, due to a cascade of signals generated
upon pollination and fertilisation that results in the senescence of
sepals/petals, but not the ovaries.
Few studies on the physiology of flowering in tropical orchids
have been conducted to date. As orchid cultivation continues to be
a highly profitable commercial market (Hew and Yong, 2004),
characterisation of tropical orchid flowering is of paramount
importance. Dendrobium crumenatum (Swartz), also known as the
pigeon orchid, is a common native epiphytic orchid species of
South-east Asia. It exhibits an interesting and unique diversion of
the normal flowering process: upon transition of the meristem
from a vegetative to a reproductive phase, floral buds develop to a
certain stage and then become ‘dormant’. These floral buds resume
growth and development after cold-induction, such as after a
heavy rainfall, and culminating into the opening of the flowers
exactly nine days after (Holttum, 1953; Corner, 1988). Full flower
opening is achieved before dawn and the flowers last for only 24 h
under natural conditions. Senescence of the flowers is indicated by
the flaccid sepals and petals. Knowledge on the physiological
processes controlling dormancy release, floral bud development,
flower opening and senescence in D. crumenatum can be applied to
commercially important flowers where it would be advantageous
to be able to control the timing of floral bud development. For
example, it would be beneficial to be able to force floral buds into
dormancy during shipment, releasing them from dormancy and to
resume normal floral bud development when required.
In this study, carbohydrates, water potential, cell wall
components and activities of cell wall-based enzymes of D.
crumenatum were analysed throughout the development of newly
induced floral buds till flower senescence. These data would allow
us a better understanding of the physiological processes, especially
the possible involvement of cell wall metabolism, in the regulation
of flowering.
2. Materials and methods
2.1. Plant material
Dendrobium crumenatum (Swartz) plants were maintained
under partially shaded conditions (PAR ranged from 100 to
250 mmol mÀ2 sÀ1; average air temperature ranged from 25 to
33 8C) in a planthouse of the Department of Biological Sciences,
National University of Singapore. All plants were watered daily,
and fertilised weekly with a foliar fertiliser (N18:P36:K18). Pots of
D. crumenatum with inducible inflorescences carrying dormant
floral buds were acclimatised at 30 8C for 24 h in temperaturecontrolled growth chambers. They were then subjected to a cold
induction at 20 8C for 24 h. Growth chambers were maintained on
a 12 h day/12 h night cycle and photosynthetic active radiation
(PAR) ranged from 10 to 20 mmol mÀ2 sÀ1. Plants of D. crumenatum
exhibit crassulacean acid metabolism, demonstrating different
carbon dioxide exchange patterns during different times of the
day. Thus, all plants were moved into the growth chambers at
1600 h, to minimise the effects of any possible temporal variations
in the plant physiology. Plants were also watered daily to minimise
dehydration stress. Floral buds or flowers for analyses of
carbohydrates, water potential, cell wall composition and cell
wall enzyme activities were selected according to their age and
features (Table 1, Fig. 3). Sepals and petals of the harvested floral
buds and flowers were separated and stored at À80 8C until use.
2.2. Carbohydrate analyses
Sepals or petals (0.1 g) were boiled in 10 ml of distilled water for
90 min. The supernatant was collected as the total soluble sugar
fraction. The residue was re-suspended in 10 ml of 10 mM sodium
Table 1
Stages of floral bud development in D. crumenatum
Features
Exposure of dormant floral buds to cold induction at 20 8C for 24 h
Green bud (ca. 1 cm long) with reddish brown tinges along ventral side,
elongation of mentum, mentum reddish brown
Light green bud (ca. 2.5 cm long), reddish brown tinges only at beginning and tip
of mentum, further elongation of mentum, length of mentum almost half
of length of whole bud
White bud (ca. 3 cm long), no splitting of sepals, elongated mentum pointing
downwards away from tip of bud
Full flower opening, sepals and petals fully expanded, lip fully protruded with
visible yellow ridges running down from midlobe to foot of column
Sepals and petals shrivelled and brownish
Timing of events is reported in relation to the time during which floral buds were released from dormancy by cold induction (denoted as day 0).
Time (day)
0
4
7
9
10
12
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Y.-M. Yap et al. / Scientia Horticulturae 119 (2008) 59–66
acetate buffer (pH 4.5) at 30 8C for 24 h. The supernatant was
collected as the fructan fraction. The remaining residue was
ground in 10 ml of 10 mM sodium acetate buffer (pH 4.5).
Amyloglucosidase (12.4 units, Rhizopus, Sigma) was added to
the ground plant materials and incubated at 45 8C for 24 h. The
suspension was centrifuged at 4000 Â g for 10 min at 4 8C. The
supernatant was collected as the starch fraction. All fractions were
kept frozen until use. Concentrations of total soluble sugars,
fructans and starch were determined by the phenol–sulphuric acid
method (Dubois et al., 1956). Glucose was used as the standard in
the assays.
2.3. Water potential analyses
Frozen sepals or petals were allowed to thaw at room
temperature (22 8C) for 20 min and were centrifuged at
10,000 Â g for 10 min at 18 8C for the collection of cell sap. Sap
osmolality was determined using a vapour pressure osmometer
(Wescor, model 5520 VAPRO). The corresponding water potential
was calculated using the Van’t-Hoff relation: water potential
(MPa) = ÀRT (mol kgÀ1), where R is the ideal gas constant
(0.00831 MPa kg molÀ1 KÀ1) and T is temperature (K) (Nobel,
1983; Maricle et al., 2007).
2.4. Light microscopy
Segments of approximately 1 cm  0.5 cm, including the
central vein, were cut about halfway from the tips of freshly
harvested sepals or petals. The fresh tissues were fixed in a
ethanol:acetic acid:formalin:water (10:1:2:7 by volume) mixture,
dehydrated in a series of increasing concentrations of ethanol,
infiltrated with melted paraffin wax, and then embedded in
paraffin wax. Embedded tissues were sectioned using a microtome
to a thickness of 10 mm. The sections were stained with toluidine
blue (0.1%) and examined using light microscopy.
2.5. Cell wall compositional analyses
Cell wall materials were prepared by using a modified method
described by Huber (1992). Frozen sepal or petal tissues were
homogenised in 95% ethanol at 4 8C and then chilled at À20 8C for
24 h. The homogenates were centrifuged at 8000 Â g for 10 min at
4 8C. Tris-buffered phenol (5 ml per gram fresh weight of sepal/
petal tissues) was added to the residue and the whole mixture was
incubated at room temperature for 45 min. The suspensions were
re-centrifuged as described above, and the residues were resuspended in 80% ethanol at À20 8C for 2 h, followed by
centrifugation again. The remaining residues were washed once
with 80% ethanol, followed by 80% acetone, and then chloroform:methanol (1:1) mixture. All organic washings were conducted at room temperature. The final residues were recovered by
filtration, washed thrice with acetone until total whitening,
yielding the crude cell wall material (ethanol-insoluble residue,
EIR). The EIR was air-dried, weighed, and then stored at À80 8C.
Cellulose content of the EIR (5 mg) was extracted following the
procedure described by Updegraff (1969) involving an initial
hydrolysis in 5 ml of acetic–nitric acid reagent (10 volumes of 80%
acetic acid:1 volume of nitric acid) at 100 8C for 30 min, followed
by a further hydrolysis in 67% sulphuric acid at room temperature
for 1 h. Total pectin in EIR (5 mg) was extracted as described by
Ahmed and Labavitch (1977), involving a complete hydrolysis in
2 ml of concentrated sulphuric acid at room temperature for
20 min.
Soluble pectins and hemicelluloses were sequentially extracted
from EIR by water (water-soluble pectin), 50 mM sodium acetate
61
buffer (pH 6.0) containing 50 mM CDTA (CDTA-soluble pectin),
50 mM sodium carbonate containing 20 mM sodium borohydride
(Na2CO3-soluble pectin), and finally 6 N sodium hydroxide
containing 0.13 mM sodium borohydride (hemicellulose). The
proportion of EIR to extractant was 1 mg:0.5 ml. All extractions
were conducted at room temperature for 2 h with constant
shaking, and then centrifuged at 10,000 Â g for 15 min at 4 8C
(O’Donoghue et al., 2002; Deng et al., 2005).
Cellulose and hemicellulose contents were assayed by the
anthrone method (Scott and Melvin, 1953). Cellulose was used as
the standard in these assays. Uronic acid contents were
determined by the m-hydroxydiphenyl method (Blumenkrantz
and Asboe-Hansen, 1973), using galacturonic acid as the standard.
2.6. Cell wall enzyme analyses
Frozen sepals or petals were ground to fine powder in liquid
nitrogen, then homogenised for 30 min in 20 mM sodium
phosphate buffer (pH 7.5) containing 1.5 M sodium chloride,
using 10 ml of extraction buffer per g fresh weight of tissues. All
enzyme extractions were performed at 4 8C. The homogenates
were centrifuged at 12,000 Â g for 20 min at 4 8C, and the
supernatants were used for the various enzymatic assays.
Pectin methylesterase (EC 3.1.1.11) activity was determined by
a continuous spectrophotometric method as described by Hagerman and Austin (1986) with some modifications. The reaction
mixture consisted of 2 ml citrus pectin solution (0.5%), 0.15 ml
bromothymol blue solution (0.01%) and 100 ml crude enzyme
extract. The pH of the solutions was adjusted to pH 7.5 each time
before assay. After adding the enzyme extract, the reaction mixture
was mixed well and allowed to stabilise for 1 min before
measuring the absorbance at 620 nm. The absorbance was again
measured after 21 min of incubation. PME activity was determined
by the difference in absorbances and was calibrated against a
galacturonic acid standard curve. One unit of PME activity
represents 1 mmol of methylester liberated per hour.
The activities of b-galactosidase (EC 3.2.1.23), b-glucosidase (EC
3.2.1.21), b-mannosidase (EC 3.2.1.25) and b-xylosidase (EC
3.2.1.37) were assayed by an adapted method described by Chin
et al. (1999). Each of the reaction mixture consisted of 0.5 ml 5 mM
p-nitrophenyl derivatives of b-D-galactopyranoside, b-D-glucopyranoside, b-D-mannopyronoside and b-D-xylopyranoside (Sigma) as
substrates respectively, 50 ml crude enzyme extract and 50 mM
sodium acetate buffer (pH 4.5) in a total volume of 2 ml. After
incubation at 37 8C for 30 min, each of the reaction was stopped by
the addition of 1 ml 0.2 M sodium carbonate, and the amount of pnitrophenol formed was determined spectrophotometrically at
415 nm. One unit of the respective glycosidase activity represents
1 mmol of p-nitrophenol released per hour. In all enzymatic assays,
boiled enzyme extracts were included as controls.
3. Results
3.1. Growth changes
Sepals and petals demonstrated similar patterns of weight
changes. Total fresh weight increased during early stages of floral
bud development and decreased upon full flower opening on day
10, due to changes in both water and dry matter contents (Fig. 1).
Increase in fresh weight: dry weight ratio observed during late
stage bud development (day 9) was due to a drastic increase in
water content, approximately 32 mg sepalÀ1 and 37 mg petalÀ1 on
day 7 to 100 mg sepalÀ1 and 97 mg petalÀ1 on day 9. A decrease in
the ratio was also observed during the senescence stage (day 12)
where water loss was much greater than dry matter loss.
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Fig. 1. Characteristics of weight changes of (A) sepals and (B) petals of D.
crumenatum at each developmental stage. Each value is the mean Æ S.E. (n = 5).
3.2. Carbohydrate and water status changes during development
During floral bud development (day 4–day 9), total soluble
sugar concentrations in both sepals and petals increased gradually,
reaching a maximum level (approximately three times that of
newly induced floral buds at day 4) upon full flower opening on day
10. Total soluble sugar concentrations subsequently decreased by
approximately 50% during senescence on day 12 (Fig. 2A). The
reserve carbohydrates, fructans and starch, decreased in concentrations throughout floral bud and flower development, up till
senescence (Fig. 2B–C). The accumulation of total soluble sugars
during flower opening was accompanied by decreases in water
potential of sepal and petal cell sap, which subsequently increased
during senescence (Fig. 2C).
3.3. Changes in anatomy during development
Cross-sections of sepals and petals from young, newly induced
floral buds (day 4) showed round parenchyma cells packed tightly
and neatly between the upper and lower epidermes (Fig. 3A). By
day 7, the parenchyma cells had enlarged and were less tightly
packed together, exhibiting some intercellular spaces (Fig. 3B).
This layer became progressively disorganised as floral buds
continued to develop and open into flowers, with larger intercellular spaces and membrane disintegration becoming apparent
(Fig. 3C and D). By senescence stage on day 12, both sepals and
petals showed almost complete absence of parenchyma cells,
although the epidermal layers remained intact (Fig. 3E).
3.4. Cell wall composition
Sepals and petals demonstrated similar patterns of changes in
cell wall composition. The amount of EIR increased from
approximately 0.2 mg (floral part)À1 to 0.8 mg (floral part)À1
throughout early floral bud development and flower opening (day
4–day 9), but decreased to approximately 0.4 mg (floral part)À1 as
flowers senesced on day 12 (Fig. 4A). Cellulose levels increased
Fig. 2. Changes in amounts of (A) total soluble sugars, (B) fructans, and (C) starch
and water potential during development of D. crumenatum flowers. Each value is
mean Æ S.E. (n = 10).
from about 5 mg (floral part)À1 to 30 mg (floral part)À1 as the floral
buds developed to maturity and decreased to about 25 mg (floral
part)À1 during senescence (Fig. 4B). Hemicellulose levels decreased
steadily from approximately 30 mg (floral part)À1 to 2 mg (floral
part)À1 throughout bud and flower development (Fig. 4C).
The amount of total pectin generally increased during floral bud
development (day 4–day 9), and remained constant (approximately 90 mg sepalÀ1 and 60 mg petalÀ1) throughout flower
opening and senescence on day 10 and day 12 respectively
(Fig. 4D). The amount of water-soluble pectin increased by about
three times during early floral bud development on day 7,
remained at these levels (approximately 6 mg sepalÀ1 and
3 mg petalÀ1) in mature buds, then decreased upon flower opening
and senescence (Fig. 4E). On the other hand, the quantity of CDTAsoluble pectin increased during bud development and remained at
high levels (approximately 20 mg sepalÀ1 and 8 mg petalÀ1) even
during senescence (Fig. 4F). The levels of Na2CO3-soluble pectin
also increased progressively during bud development, remaining
at high levels (approximately 60 mg sepalÀ1 and 40 mg petalÀ1)
during flower opening and then decreased during senescence
(Fig. 4G). Maximum amounts of water-soluble pectin, CDTAsoluble pectin and Na2CO3-soluble pectin were all observed during
late floral bud stage at day 9, indicating a maximum level of pectin
solubility during this developmental period. The amount of
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63
4. Discussion
Fig. 3. Changes in anatomical features of D. crumenatum during flower
development. (I) Developmental stages of floral buds/flowers: A, day 4; B, day 7;
C, day 9; D, day 10; E, day 12 floral buds/flowers. Transverse sections of (II) sepals
and (III) petals corresponding to those in I were stained with toluidine blue. Arrows
represent areas with parenchyma cell disruption and/or intercellular spaces.
insoluble pectin decreased during floral bud development (day 4–
day 9), and subsequently increased upon flower opening up till
senescence (Fig. 4H).
3.5. Activities of wall-based enzymes
Colorimetric analyses using fresh tissue extracts demonstrated
the extracts did not contain components to hydrolyse carboxymethylcellulose, filter paper and polygalacturonic acid. As such,
we concluded that sepals and petals of D. crumenatum had no or
very insignificant levels of cellulase and polygalacturonase
activities.
Pectin methylesterase activity increased during early floral bud
development and decreased drastically during late floral bud
development, and remained at low levels throughout flower
opening and senescence stages (Fig. 5A). Maximum activities of bgalactosidase, b-glucosidase, b-mannosidase and b-xylosidase
were observed during floral bud development (day 7 or day 9,
Fig. 5B–E). Of all the enzymes tested, the major ones were bglucosidase [0.3–5 units (floral part)À1], pectin methylesterase
[0.5–4.3units (floral part)À1] and b-galactosidase [0.4–0.7 units
(floral part)À1]. The activities of b-mannosidase and b-xylosidase
were lower [0.04–0.2 units (floral part)À1]. With the exception of
b-galactosidase, activities of all other tested glycosidases exhibited significant decreases during flower senescence (Fig. 5B–E). bgalactosidase activity decreased slightly during flower opening,
and then increased again upon senescence.
There is great variation among flowering plants in the
physiology regulating floral bud development, flower opening
and senescence (O’Donoghue, 2006). The results obtained from the
present study provided novel insights into the floral bud
development and flower opening processes in tropical orchids.
Flower opening and senescence in D. crumenatum were found to
be dependent on carbohydrate metabolism and water relations
(Fig. 2). The high accumulation of reserve carbohydrates in sepals
and petals of D. crumenatum during early stages of bud
development was not unexpected as inflorescences and flowers
are considered to be sinks for assimilates (Hew and Yong, 2004).
During the flowering process in orchids, the priority of assimilate
partitioning usually follows this decreasing order: inflorescences ) young leaves ) shoots (Hew and Yong, 2004). Prior to the
onset of flower opening in D. crumenatum, the significant breakdown of reserve carbohydrates and increase in levels of soluble
sugars resulted in a more negative water potential. This resulted in
a larger water potential gradient, possibly causing a greater water
influx into the expanding cells and subsequent turgor changes,
which had been suggested to be a major force in driving flower
opening (van Doorn and van Meeteren, 2003). Flower opening
regulated by changes in carbohydrate levels and water relations
has also been reported in species such as carnation (Acock and
Nichols, 1979), daylily (Bieleski, 1993) and creeping bellflower
(Vergauwen et al., 2000). Contradictory to our results, however,
Sonia roses did not show a relationship between carbohydrate and
water status during flower opening (Evans and Reid, 1988). During
rapid petal expansion in roses, starch hydrolysis occurred, but
osmotic potential of the cell sap became less negative, reducing the
water potential gradient (Evans and Reid, 1988). Hence, it was
proposed that mobilisation of storage carbohydrates and water
uptake were not the main controlling factors in rose petal growth,
but rather other physiological factors such as cell wall extensibility
might be involved (Evans and Reid, 1988).
Cell wall metabolism in flowers has been studied in great detail
in only two other species—sandersonia and carnation (de Vetten
and Huber, 1990; de Vetten et al., 1991; O’Donoghue et al., 2002),
but cell wall alterations were not investigated in young developing
floral buds in these studies. Studies conducted on daylily focused
on activities of cell wall hydrolases from young floral buds till
senesced flowers, but no accompanying analyses on cell wall
compositional changes were reported (Panavas and Rubinstein,
1998; Panavas et al., 1998). With these in mind, and the proposals
that cell wall metabolism might be involved during flower opening
and senescence (O’Donoghue, 2006), we examined the changes in
anatomy of sepals and petals of D. crumenatum during floral bud
and flower development. The increasing disorganisation of
parenchyma cells and appearance of intercellular spaces observed
(Fig. 3) during the development of D. crumenatum floral buds
resembled those observed during the opening of sandersonia and
daylily flowers (Panavas et al., 1998; O’Donoghue et al., 2002),
indicating alterations in cell wall constituents.
In our study, cell wall composition and activities of cell wallbased enzymes were expressed as ‘per floral part’ basis and similar
trends were obtained when expressed as ‘per flower basis’ (data
not shown). The accumulation of cell wall cellulose during floral
bud development and subsequent breakdown during senescence
in D. crumenatum (Fig. 4A) were similar to those of carnation and
sandersonia (de Vetten and Huber, 1990; O’Donoghue et al., 2002).
However, while both D. crumenatum and sandersonia flowers
showed no cellulase activity, the enzyme was detected in daylily
flowers, indicating possible hydrolysis of cellulose by cellulase in
daylily (Panavas et al., 1998). Hemicellulose content decreased
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Fig. 4. D. crumenatum sepal and petal cell wall components, expressed on a per floral part (sepal or petal) basis, at each developmental stage: (A) EIR, (B) cellulose, (C)
hemicellulose, (D) pectin, (E) water-soluble pectin, (F) CDTA-soluble pectin, (G) Na2CO3-soluble pectin, and (H) insoluble pectin. Each value is mean Æ S.E. (n = 5).
steadily in D. crumenatum, while that in sandersonia showed no
apparent changes (O’Donoghue et al., 2002). The breakdown of
hemicelluloses in D. crumenatum resembled the changes observed
during the ripening of fruits such as grapes (Deng et al., 2005) and
carambola (Chin et al., 1999). Hemicelluloses (or glycans) play
important structural roles in the cross-linking of cellulose in the
cell walls and the breakdown of hemicelluloses may contribute to
the alterations in primary cell wall structure (Brummell, 2006).
Activities of various glycosidases were detected in D. crumenatum
and were observed to alter with the development of the floral buds
(Fig. 5C–E). Various glycosidases had also been reported in
carnation and were suggested to be involved in the hydrolysis
of hemicelluloses (de Vetten et al., 1991). Glycosidases had also
been reported in fruits such as capsicum (Sethu et al., 1996) and
carambola (Chin et al., 1999). The family of glycosidases is known
to play a crucial role in the degradation of various cell wall
polysaccharides, allowing the remodelling of the cell wall structure
(Minic, 2008).
Pectin hydrolysis and polygalacturonase activity were not
detected in D. crumenatum flowers (Fig. 5). These were similar to
those observed in sandersonia flowers (O’Donoghue et al., 2002).
However, daylily flowers were reported to exhibit increasing
polygalaturonase activity in senescing flowers, indicating the
possibility of pectin hydrolysis in these flowers (Panavas et al.,
1998). While pectin synthesis or accumulation occurred in D.
crumenatum, there was also considerable pectin solubilisation.
Water-soluble and CDTA-soluble pectins are relatively weakly
bound to cell wall polysaccharides by molecular entanglements,
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Y.-M. Yap et al. / Scientia Horticulturae 119 (2008) 59–66
Fig. 5. Changes in activities, expressed on a per floral part (sepal or petal) basis, of
(A) pectin methylesterase, (B) b-galactosidase, (C) b-glucosidase, (D) bmannosidase and (E) b-xylosidase during development of D. crumenatum
flowers. Each value is mean Æ S.E. (n = 5).
hydrophobic forces, weak ionic bonds or ionic calcium bridges,
while Na2CO3-soluble pectins are more strongly attached to the
cell wall via covalent bonds (Brummell, 2006). In D. crumenatum,
the types of bonds and bond strengths that interlink pectin
molecules appeared to alter as the floral buds and flowers
continued to develop. There were increasing proportions of pectins
that were susceptible to the solubilising agents during floral bud
development, and decreased pectin solubility during flower
opening and senescence (Fig. 4E–H). Pectin solubilisation is one
65
ripening-associated cell wall modification that has been widely
reported in an array of fruits, and different species of fruits also
exhibited variations in pectin modifications (Chin et al., 1999;
Brummell and Harpster, 2001; Deng et al., 2005; Vicente et al.,
2007). Cell wall swelling, which would enhance access of
hydrolytic enzymes to their substrates and promote polymer
disassembling, had been correlated to increased pectin solubilisation (Redgwell et al., 1997).
Pectin methylesterase, which de-esterifies pectins, had been
suggested to increase pectin solubilisation by creating electrostatic
repulsion between negatively charged molecules that could result
in the loosening of weakly attached pectins from the cell wall
(Grignon and Sentenac, 1991). In D. crumenatum, the largest
increase in pectin solubilisation, which was observed during floral
bud development, coincided with maximum pectin methylesterase activity (Fig. 5A). However, the significant decrease in pectin
methylesterase activity, shortly after the peak, was not accompanied by a drastic decrease in pectin solubility (Fig. 5E–H). In
sandersonia, pectin methylesterase activity (units flowerÀ1 basis)
peaked upon mature flower stage, then decreased till senescence
(O’Donoghue et al., 2002). In daylily, pectin methylesterase activity
(units mg proteinÀ1 basis) decreased from young floral buds stage
till flower opening, similar to D. crumenatum (units mg proteinÀ1
basis, data not shown). Such variations among the different species
may be the result of, or cause(s) for, the differences in timeframes
of floral bud development, flower opening, senescence and
longevity of the flowers (O’Donoghue, 2006). Different species of
fruits also demonstrated variations in cell wall pectic modifications during ripening (Brummell, 2006).
There are many other factors that may also regulate pectin
solubilisation and other polymer modifications. One such candidate could be b-galactosidase, an enzyme responsible for the
degradation of cell wall b-galactan, regulating cell wall flexibility,
intercellular connections and cell wall porosity, and affecting
mobility of enzymes within the cell wall matrix (Brummell and
Harpster, 2001; Brummell, 2006). In vitro treatments of cell wall
preparations from papaya with b-galactosidase resulted in
increased pectin solubilisation (Ali et al., 1998). In D. crumenatum,
the activity of b-galactosidase remained relatively constant during
late floral bud and flower opening stages, and increased slightly
during senescence (Fig. 5B). This might account for the maintenance of CDTA-soluble pectins (Fig. 4F). In sandersonia, bgalactosidase activity (units flowerÀ1 basis) increased during late
floral bud up till wilted flower stage, and decreased significantly
during senescence (O’Donoghue et al., 2002). On the other hand, bgalactosidase activity (units mg proteinÀ1) in daylily was constant
throughout bud development and flower opening, and only
increased during wilting and senescence stages, similar to that
observed in D. crumenatum (units mg proteinÀ1, data not shown). It
is possible that non-enzymic mechanisms might also be involved
in cell wall modifications. In tomato, ascorbate-generated hydroxyl radicals demonstrated non-enzymic scission of cell wall
polysaccharides, and caused an increase in pectin solubilisation
(Dumville and Fry, 2003). Cell wall modification is thus a complex
process, and it would be worthwhile to identify other non-enzymic
factors, and also investigate the possible interactions between the
various enzymic and non-enzymic mechanisms involved in the
process.
There are questions on whether flowering is part of a
senescence programme that has already started (O’Donoghue
et al., 2002). Carbohydrate and cell wall polymer alterations are
considered to be events associated with cell death and senescence
(Rubinstein, 2000). Previously, studies on carnation and sandersonia suggested that just prior to flower opening (late floral bud
stage), a certain degree of cell wall dismantling had already started
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Y.-M. Yap et al. / Scientia Horticulturae 119 (2008) 59–66
(de Vetten and Huber, 1990; de Vetten et al., 1991; O’Donoghue
et al., 2002). Our results showed that senescence-related events
such as cell wall disassembly occurred as early as during young
floral bud stage (day 7), suggesting the early onset of a senescence
programme during floral bud development. Flower opening, thus,
appears to be a continuum of a senescence programme that had
already commenced. It remains to be tested if inhibitors of cell wall
hydrolases and/or senescence-inhibiting chemicals could delay
floral bud development and flower opening. Such techniques
would be beneficial to the horticultural industry that commonly
depends on refrigeration to delay flower opening; the techniques
could also help lower energy consumption.
In conclusion, floral bud development, flower opening and
flower senescence in D. crumenatum were tightly regulated by
changes in carbohydrate and water status, and cell wall
metabolism. Senescence-indicative events such as cell wall
disassembling were found to occur during early floral bud
development, suggesting the early onset of a senescence programme. Cell wall metabolism is a common feature among D.
crumenatum, sandersonia, carnation and daylily but the timing and
manner of modifications vary between the species. Whether these
differences are the cause for the diversity in the rates of floral bud
development, time of flower opening, and longevity of the flowers
in the various plants remain to be investigated. Investigating the
possible roles of non-enzymic mechanisms in controlling cell wall
disassembly could provide further insight to the involvement of
cell wall metabolism in controlling flowering, and the relation
between flowering and the in-built senescence programme of the
plant.
Acknowledgements
We thank H.H. Yeoh for providing the various enzymatic
substrates, P.L. Chong and B.L. Tay for their technical help in the
experiments, and T.K. Ong, W.P. Yap, H.Q. Lim and S.L. Ng for their
assistance in many ways. We also thank the National University of
Singapore for sponsoring the Research Scholarship for Y.M. Yap.
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[...]... variation in cell wall metabolism that regulates flowering (Panavas et al 1998; O’Donoghue et al 2002) In the above-mentioned studies on the possible involvement of cell wall modifications and cell wall remodelling in regulating flowering, cell wall changes were compared only between stages of mature bud (just prior to opening), opening flower, mature flower, wilting flower and senesced flower (de... (Chin et al 1999; Deng et al 2005) In daylily flowers, analyses of cell wall composition were not published, but reported changes in activities of cell wall- based enzymes during flower development suggested the involvement of cell wall metabolism in flowering (Panavas et al 1998) While cellulase activity was detected in daylily flowers, it was reported to be absent in sandersonia flowers, indicating... between the flowers shows certain similarities and differences in cell wall changes associated with flowering In carnation flowers, full flower opening was accompanied by increases in contents of cell wall cellulose, total pectins, chelator-soluble pectins, carbonate-soluble pectins and neutral sugars Upon senescence of the flowers, all of these cell wall components, except chelator-soluble pectins, decreased... the cells cease growth (Cosgrove 1999) The dynamic structure of the cell wall is important in regulating cell expansion and cell growth For example, cell wall loosening is a pre-requisite for the incorporation of newly synthesized wall polymers during cell expansion and cell growth (Carpita and McCann 2000) Plant cell walls also have important roles in controlling fruit ripening The degree of cell wall. .. (Rubinstein 2000; Wagstaff et al 2003) In Ipomoea, dynamic structural changes such as cell enlargement, modification of cell shape and reduction in cell wall thickness occurred in the inner epidermal cells even before flower opening (Phillips and Kende 1980) In sandersonia, intercellular air spaces and increasingly disorganized packing of parenchyma cells also occurred prior to flower opening (O’Donoghue... that may involve analysing and comparing cell wall modifications between floral buds showing normal and abnormal opening patterns 3 Chapter 2 Literature Review 2.1 Plant cell wall Each plant cell comprises a specialised and complex cell wall that serves many functions (Cosgrove 1999) The cell walls provide structural support and maintain the shape of the cells, act as a protective barrier against water... published regarding cellulose degradation As the main component of the cell wall matrix, cellulose is expected to undergo distinct alterations during cell wall changes (Fischer and Bennett 1991; Rose and Bennett 1999) Cell wall cellulose content decreased during ripening of grapes (Deng et al 2005) However, during the ripening of pear (Pyrus communis), tomato and avocado, cellulose levels remained constant... hemicelluloses held tightly by hydrogen bonds (Brummell 2006) 7 2.2 Cell wall metabolism Various regulated cell wall architecture changes occur with the development of plants (Carpita and McCann 2000) Fruit ripening is one developmental event whereby many changes occur in the cell walls, resulting in the final texture of the fruit (Brummell 2006) Consequently, the majority of the information on cell wall. .. Pectin solubilisation One of the major changes in cell wall pectins with the development of plants is the increasing solubilisation of pectins, as commonly observed during the ripening of fruits (Brummell and Harpster 2001) Pectin solubilisation is usually measured as the increase in ease of extractability of pectins by various extractants, and can be extrapolated to bond changes within the cell wall. .. question on whether flowering could be a process regulated by the senescence programme of the plant Some supporting evidence include the increase in oxidation of membrane components prior to flower opening in daylily (Panavas and Rubinstein 1998), and the increasing trend of DNA laddering throughout petal development in alstroemeria (Wagstaff et al 2003) Studies on carnation and sandersonia 1 flowers demonstrated ... the flowers shows certain similarities and differences in cell wall changes associated with flowering In carnation flowers, full flower opening was accompanied by increases in contents of cell wall. .. changes in activities of cell wall- based enzymes during flower development suggested the involvement of cell wall metabolism in flowering (Panavas et al 1998) While cellulase activity was detected in. .. 1998; O’Donoghue et al 2002) In the above-mentioned studies on the possible involvement of cell wall modifications and cell wall remodelling in regulating flowering, cell wall changes were compared