THE PHOTOPROTECTIVE ROLE OF ANTHOCYANIN PIGMENTS
IN LEAF TISSUES
By
NICOLE M. HUGHES
A Dissertation Submitted to the Graduate Faculty of
WAKE FOREST UNIVERSITY
GRADUATE SCHOOL OF ARTS AND SCIENCES
In Partial Fulfillment of the Requirements
for the Degree of
DOCTOR OF PHILOSOPHY
in the Department of Biology
May 2009
Winston-Salem, NC
Approved by:
William K. Smith, Ph.D., Advisor
Examining Committee:
Gloria K. Muday, Ph.D.
Peter D. Weigl, Ph.D.
Kathleen A. Kron, Ph.D.
Taylor S. Feild, Ph.D.
ACKNOWLEDGEMENTS
I would first like to thank my parents, Randy and Theresa, for their love and support in
everything I’ve ever done in my life. At the interface between family and science, I
thank my science fathers—Bill Smith and Howie Neufeld, for their guidance, advice, and
support. I also thank my fellow lab mate, good friend, and mentor, Dan Johnson. I don’t
thank Keith Reinhardt for anything, but I acknowledge that he has served some purpose
in my life here as my scientific and personal antagonist for three years. Thanks also to
my dearest friends—Alison Northup, Ali Farr, Adriana Sanchez, Ellen and Adam
Koehler, Bruce Moser, Catherine Reinhardt, Sara Venables, Lauren Eiter, and Sarah
Higgins.
Chapter-specific acknowledgements
Chapter 1- I thank Drs. L. Donovan and H. Neufeld for use of the spectroradiometric
equipment, Dr. A. McCauley and the microscopy facility at WFU, C. Machalaba and S.
Bissett for technical assistance, and the anonymous reviewers of this work, whose helpful
suggestions greatly improved the quality of this manuscript. This work was supported
financially by the graduate school at Wake Forest University.
Chapter 2- Craig Brodersen and Nate Poirier (University of Vermont) provided technical
assistance with fluorescence profiling, Dr. Howard Neufeld (Appalachian State
ii
University) provided use of optical filters and the Li-1800, and Tim Anderson
(PalmHammock Orchid Estate) graciously donated plant materials.
Chapter 3- I would like to thank Spencer Bissett for technical assistance and Dr. Anita
McCauley of the microscopy facility at WFU for help with microscopic imaging. This
study was supported financially by Wake Forest University and a Research Experience
for K-12 Teachers (RET) grant to CBM (National Science Foundation).
Chapter 4- Spencer Bissett and Kelsey McDowell were instrumental in their help in the
field and lab, respectively. Funding for this project was provided by the Vecellio Fund at
Wake Forest University.
iii
TABLE OF CONTENTS
ACKNOWLEDGEMENTS……………………………………………………………….ii
LIST OF TABLES……………………………………………………………………….vii
LIST OF FIGURES……………………………………………………………………..viii
ABSTRACT……………………………………………………………………………….x
CHAPTER I
Anthocyanin pigments: An introduction………………...………………………..1
CHAPTER II
Attenuation of incident light in Galax urceolata (Diapensiaceae): Concerted
influence of adaxial and abaxial anthocyanic layers on photoprotection.
Published in American Journal of Botany (2007)
ABSTRACT……………………………………………………………...25
INTRODUCTION…………….…………………………………………26
MATERIALS AND METHODS………………………………………...29
RESULTS……………………………………………………..…………33
DISCUSSION……………………………………………………………35
LITERATURE CITED………………………………………………..…39
CHAPTER III
Optical effects of abaxial anthocyanin on absorption of red wavelengths by
understory species: revisiting the back-scatter hypothesis.
Published in Journal of Experimental Botany (2008)
iv
ABSTRACT……………………………………………………………...51
INTRODUCTION……………………………………………………….52
MATERIALS AND METHODS………………………………………...55
RESULTS & DISCUSSION…………………………………………….59
LITERATURE CITED…………………………………………………..65
CHAPTER IV
Blackwell Publishing Ltd
Coordination of anthocyanin decline and photosynthetic maturation in juvenile
leaves of three deciduous tree species.
Published in New Phytologist (2007)
ABSTRACT…………………………………………………………...…82
INTRODUCTION…………………………………………………….…83
MATERIALS AND METHODS……………………………….……..…86
RESULTS…………………………………………………………….….91
DISCUSSION……………………………………………………………96
LITERATURE CITED…………………………………………………103
CHAPTER V
Seasonal photosynthesis and anthocyanin production in ten broadleaf evergreen
species.
Published in Functional Plant Biology (2008)
ABSTRACT…………………………………………………………….118
INTRODUCTION…………………………………………………...…119
MATERIALS AND METHODS……………………………………….122
RESULTS………………………………………………………………127
v
DISCUSSION………………………………………………………..…130
LITERATURE CITED…………………………………………………133
CHAPTER VI
Conclusions and Future Directions……………………………………………..149
CURRICULUM VITAE……………………………………………………………..…153
vi
LIST OF TABLES
Table 2.1. Pigment content and leaf thickness values for the four leaf types of Galax
urceolata used in abaxial/adaxial optical experiments…………………….…...43
Table 3.1. Chlorophyll data for abaxially anthocyanic (red) and acyanic (green) portions
of Begonia heracleifolia leaves…………………………………………………80
Table 5.1. Detailed descriptions of five red and five green broadleaf evergreen
species ……………………………………………………………...…………..137
Table 5.2. Winter photosynthetic and non-photosynthetic pigment content of red and
green broadleaf evergreen species………………………………………..…….139
vii
LIST OF FIGURES
Figure 1.1. Anthocyanins depicted during all life stages and seasons…………..………17
Figure 1.2. The biosynthetic pathway for anthocyanin………………………….………19
Figure 1.3. Leaves exhibiting light-dependence of anthocyanin synthesis…………..…21
Figure 1.4. Anatomical location of anthocyanins………………………………….……23
Figure 2.1. Cross sections of Galax urceolata leaves exhibiting the four combinations of
pigmented surfaces seen in winter leaves………………..………………………45
Figure 2.2. Spectral scans of Galax urceolata leaves exhibiting four combinations of
pigmented surfaces seen in winter leaves………………………………………..47
Figure 2.3. Recovery of maximum PSII efficiency (Fv/Fm) for abaxial and adaxial
surfaces of red and green Galax urceolata leaves……………...………………..49
Figure 3.1. Adaxial view of Begonia heracleifolia leaf, and chlorophyll fluorescence
profiles in red and green variegated leaf sections under red, green, and
blue light……………………………………………………………………...….70
Figure 3.2. Spectral distribution of ambientlight, the red optical filter used in
experiments, and light reflected by red abaxial surfaces B. heracleifolia ………72
Figure 3.3. Optical properties (absorptance, transmittance, and reflectance) of adaxial
and abaxial surfaces of anthocyanic and acyanic tissues of B. heracleifolia…....74
Figure 3.4. Quantification of chlorophyll fluorescence profiles according to leaf depth
for abaxially anthocyanic and acyanic B. heracleifolia tissues under red, blue, and
green light………………………………………………………………………..76
viii
Figure 3.5. Photosynthetic light response curves of abaxially acyanic/green and
anthocyanic/red leaf tissues under red light……………………………...………78
Figure 4.1. Whole branches and microscopic sections of leaves from nodes 1-5 of
L. styraciflua, A. rubrum, and C. canadensis …………………………...……...108
Figure 4.2. Pigment concentrations and leaf thicknesses expressed as a percent of mature
values for L. styraciflua, A. rubrum, and C. canadensis…………………….….110
Figure 4.3. Percent maximum photosynthesis (Asat) and leaf conductance of water vapor
(g) values by leaf node, expressed as a percent of mature-leaf values, for L.
styraciflua, A. rubrum, and C. canadensis…………...…………………………112
Figure 4.4. Tip-down (nodes 1-5) chlorophyll a/b ratios for species with green
developing leaves and red developing leaves……..............................................114
Figure 4.5. The relationships between photosynthesis maturation and corresponding
changes in chlorophyll per unit area, leaf thickness, and anthocyanin for L.
styraciflua, A. rubrum, and C. canadensis………………………………...……116
Figure 5.1. Winter maximum light capture efficiency of PSII (Fv/Fm), light-saturated
photosynthesis (Asat), and leaf conductance (g) for red and green broadleaf
evergreen species measured across a range of ambient temperatures…………141
Figure 5.2. Cold, mild, and warm winter day measurements of Asat, g, and Fv/Fm for
green-leaf and red-leaf species…………………………………………………143
Figure 5.3. Seasonal light response curves for red and green broadleaf evergreen
species…………………………………………………………………………..145
Figure 5.4. Percent decrease in maximum observed winter Asat relative to summer values
for red and green broadleaf evergreen species…………………………………147
ix
ABSTRACT
Hughes, Nicole M.
THE PHOTOPROTECTIVE ROLE OF ANTHOCYANIN PIGMENTS
IN LEAF TISSUES
Dissertation under the direction of William K. Smith, Ph.D.,
Charles H. Babcock Professor of Botany
Anthocyanins are vacuolar pigments most commonly responsible for red to purple
coloration in plant tissues. Because they absorb strongly in the blue-green waveband,
anthocyanins effectively reduce internal light within the leaf, which may be beneficial
under high-light stress conditions. However, pigment patterns in natural systems often
appear inconsistent with a photoprotective function (e.g. anthocyanins in abaxial leaf
surfaces), and their absence in many species exposed to high-light stress suggests that
anthocyanins may not be necessary for photoprotection. Furthermore, the presumed
dynamic interaction between anthocyanin synthesis and relative need for photoprotection
has yet to be quantitatively described. These issues, among others, have stalled acceptance
of a photoprotective function of anthocyanin pigments, and are addressed in the studies
presented here.
Measurements of leaf optics and chlorophyll fluorescence were used to compare
optical and photosynthetic effects of abaxial anthocyanins in vivo. In a species where
abaxial leaf surfaces are naturally exposed to high light stress in the field (Galax urceolata),
anthocyanins appeared to function similarly to anthocyanins in adaxial surfaces—absorbing
x
strongly in the blue-green wavelengths, and reducing high-light stress (i.e. photoinhibition
of photosynthesis). In a shade-adapted, abaxially-red understory species (Begonia
heracleifolia), a photoprotective function was also supported, but through attenuation of
internally-scattered green light transmitted through the upper leaf surface. This function
may be adaptive during periodic exposure to high intensity sunflecks or sun-patches, which
are potentially damaging to shade-adapted plants.
In developing leaves of three deciduous-tree species, anthocyanin disappearance
corresponded with development of ~50% mature photopigment concentrations, ~80%
lamina thickness, and differentiation of the mesophyll into palisade and spongy layers.
This conserved pattern in anthocyanin loss during development between species is
consistent with a controlled coupling of anthocyanin concentration and relative need for
photoprotection.
Relative photosynthetic capacity was compared for five species exhibiting
anthocyanin production in winter leaves and five species lacking anthocyanin. It was
expected that species with red winter leaves would correspond with those exhibiting
diminished photosynthetic capacity, rendering them in greater need for photoprotection.
This hypothesis was not supported, however, and the reason why some species require
anthocyanin pigments while others do not during winter remains unknown.
In conclusion, these studies are generally consistent with a photoprotective function
of anthocyanins in leaf tissues.
xi
CHAPTER I
ANTHOCYANIN PIGMENTS: AN INTRODUCTION
1
Anthocyanins are vacuolar, flavonoid pigments most commonly responsible for red,
purple, and blue coloration in plant tissues. They are most often associated with coloration
in flowers and fruits, though anthocyanins may be synthesized in virtually all plant tissues,
and observed in leaves during all ontogenetic stages and seasons (Figure 1). Though the
evolutionary history of anthocyanin has yet to be fully characterized, it is known that the
pigments are absent in algae, present in some bryophytes, ferns, and conifers, and are
virtually ubiquitous in angiosperms (Lee and Gould 2002). Despite their widespread
distribution, however, the functional significance of anthocyanin pigments remains
unresolved (see reviews by Chalker-Scott 1999, Manetas 2006, Ougham et al. 2008).
Furthermore, factors which dictate their presence in certain ontogenetic stages, tissues, and
species, but not others, are not yet fully understood (Hughes and Smith 2007). The
objective of this dissertation was to address these unresolved issues in anthocyanin research
using naturally anthocyanic and acyanic plant systems and current ecophysiological
techniques.
Biochemistry of anthocyanin synthesis
Anthocyanins are secondary metabolites derived from the flavonoid branch of the
phenylpropanoid pathway. Flavonoids are derived from the precursors malonyl-CoA and
p-coumaroyl-CoA, and the biochemical pathway of anthocyanin synthesis from these
precursors is illustrated in Figure 2 (derived from Holton and Cornish 1995). Regulatory
mechanisms for flavonoid synthesis are primarily at the level of transcriptional control for
the structural genes involved in each step of the pathway (Davies and Schwinn 2003).
2
Anthocyanin gene expression is almost always dependent on light (Mancinelli
1993; Figure 3A), the intensity of which is also directly proportional to anthocyanin
concentration (Figure 3B; Mancinelli 1993; Hughes et al. 2005). Genetic expression
occurs under two distinct light conditions, involving two different processes and groups of
photoreceptors. Under low-fluence conditions (i.e. light intensities typical of the
understory), anthocyanin production is significant, but generally low in quantity, and shows
the general characteristics of a low fluence response (LFR) of phytochrome B (e.g. an
action spectrum with only one peak in the red wavelengths, activation is reversible under
far-red light and follows the Bunsen-Roscoe law of reciprocity) (Mancinelli 1993).
Anthocyanins may be induced in greater quantities under prolonged exposure to high
irradiance as a high-irradiance response (HIR). This process shows peaks in the action
spectra not only in the red, but also UV-A, UV-B and blue wavelengths, indicating
involvement of phytochrome A (activated by red light), cryptochrome (blue and UV-A),
and another, currently unknown, UV-B receptor (Mancinelli, 1993). Under high light,
phytochrome and cryptochrome are synergistic, and anthocyanin synthesis under exposure
to red + blue/UV-A is greater than the sum of anthocyanin production under red light alone
and blue/UV-A light alone. Once these sensory pigments are activated, they themselves
activate (i.e. phosphorylate or reduce) transcription factors involved in expression of the
anthocyanin structural genes.
However, high light alone is not sufficient to induce anthocyanin synthesis. Clearly,
not every plant growing in full sunlight is red, and many leaves growing under high light
will turn red only under certain growth conditions, usually those associated with abiotic
stress. More specifically, heat/cold, drought, osmotic, and nutrient stress are known to
3
induce anthocyanin synthesis in many species under high light (see Chalker-Scott 1999 for
review), as will the onset of certain ontogenetic stages or seasons which render plants
vulnerable to these stresses. The general requisite of stress in addition to light suggests that
additional regulatory mechanisms are in place to control anthocyanin synthesis in addition
to those which are light-mediated.
Hydrogen peroxide (H2O2) is known to function as a regulatory molecule in stress
perception and signal transduction, including anthocyanin synthesis (Vanderauwera et al.
2005). Indeed, many, if not all, of the conditions described above also correspond with
elevated production of radical oxygen species (ROS) and hydrogen peroxide. As expected,
the H2O2-inducible transcriptional cluster involved in anthocyanin synthesis is distinct from
the cluster inducible by light (Vanderauwera et al. 2005), though the interplay between
these two systems is not yet clear. In addition to hydrogen peroxide, sugars also play a role
in stress signaling in plant systems (Smeekens 2000; Rolland et al. 2002; Rook and Bevan
2003), and elevated levels of reducing sugars are known to induce anthocyanin synthesis
(e.g. Weiss 2000, Teng et al. 2005; Hughes et al. 2005; Murakami et al. 2008); however,
the protein(s) involved in sugar sensing, and the downstream effects on anthocyanin
synthesis, have not yet been fully characterized, though several signaling intermediates
have been identified (Vitrac et al. 2000; Teng et al. 2005). In a general ecophysiological
context, it is possible that stressful conditions which require anthocyanin may also
correspond with those that have high sugar concentrations and/or elevated production of
H2O2, which may act as a natural signal inducing anthocyanin gene expression (Teng et al.
2005; Hughes et al. 2005).
4
Synthesis of anthocyanins occurs in the cytoplasm until the precursor anthocyanidin
(last row of molecules in Figure 2). Up to 19 different naturally-occurring anthocyanidins
have been reported in plant tissues (Iwashina 2000), though the bronze-colored cyanidin-3glucoside is the most common form encountered in leaves (Lee and Gould, 2002). In the
cytoplasm, anthocyanidin is tagged with a glutathione by glutathione-S transferase, and is
transported via glutathione pump into the vacuole (Marrs et al. 1995). It is within the
vacuole (Figure 4) that anthocyanins take on their species-specific coloration, which results
from hydroxylation, methylation, sugar addition, or acylation.
Physiological function of anthocyanins in plant tissues
From within the vacuole, anthocyanins may function physiologically in many ways.
Because they absorb strongly in the blue-green waveband, anthocyanins effectively reduce
internal light within the leaf. It is important to note, however, that because these pigments
are spatially separated from the photosynthetic pigments, light-energy absorbed by
anthocyanins is not used for photosynthesis, but is rather dissipated as heat. The remaining,
un-absorbed wavelengths (i.e. those reflected or transmitted) will continue to be transmitted
through leaf cells, where they may or may not be absorbed by photosynthetic pigments
(Gould 2004). The optical properties of anthocyanins are therefore of functional
importance, as they directly influence the wavelengths and intensity of light available to be
used for photosynthesis within a leaf. Pietrini et al. (2002) estimated that anthocyanins in
Zea mays could intercept up to 43% incoming photosynthetically active radiation (PAR),
primarily between 400-600 nm, which reduced apparent quantum yield of CO2 fixation and
non-cyclic electron transport of PSII proportionately. Considering that blue-green light
5
contains the most highly-abundant, high-energy wavelengths of the solar spectrum, that
blue-green light deeply penetrates leaf tissues (Cui et al. 1991), and that wavelengths
responsible for photoinhibition and photo-bleaching are primarily between 520 and 680 nm
(Merzlyak and Chivkunova 2000), the value of blue-green light absorbing pigments during
high sunlight exposure becomes increasingly apparent. The down-regulation of internal
light for photoprotection has thus become a popular explanation for the widespread
occurrence of anthocyanin pigments in leaves and other photosynthetic tissues, especially
in high-light habitats. In vivo, red tissues exhibit less photoinhibition of photosynthesis
under blue and green light than green tissues of the same species, but equal photoinhibition
under red light, which is poorly absorbed by red pigments (Smillie and Hetherington 1999;
Feild et al. 2001; Hughes et al. 2005). Red leaves also appear to exhibit shade
characteristics relative to green conspecifics (Gould et al. 2002; Manetas et al. 2003;
Hughes and Smith 2007; Kyparissis et al. 2007). However, other studies have found no
evidence for a photoprotective function (e.g. Burger and Edwards 1996; Dodd et al. 1998;
Lee et al. 2003; Kyparissis et al. 2007).
Because anthocyanins increase energy absorption by the leaf, and anthocyanins
convert light energy into heat, it has also been proposed that anthocyanins may function to
warm leaves and increase rates of transpiration and metabolism, or protect against cold
temperatures (Smith 1909). This would be most effective in colder climates, as leaf energy
balance models dictate that leaves may be more easily uncoupled from air temperature
when air temperature is low (Campbell and Norman 1998). Indeed, some evidence exists
for a warming function of anthocyanin in other plant tissues (i.e. cones) at high altitudes
(Sturgeon and Mitton, 1980). In the case of leaves, however, there is much evidence
6
against a warming function of anthocyanins in tropical and temperate climates (e.g. Lee et
al. 1979; Lee et al. 1987; Gould et al. 1995; Lee et al. 2003), though leaf temperatures have
not been compared under very cold temperatures. Also inconsistent with a warming
function of anthocyanin is the tendency for leaves on the outer (most sunlit) portions of the
crown to synthesize anthocyanins, as these are the leaves that are exposed to the greatest
windflow, and thus, the most coupled with air temperature within the canopy (Campbell
and Norman 1998).
In addition to optical functions of anthocyanins, the pigment has also been
implicated as functioning biochemically as an in vivo antioxidant. There has been some
debate, however, as to how effective a vacuolar antioxidant could be in protecting tissues
against radical oxygen species (ROS) generated in the chloroplast (Manetas 2006). It has
recently been shown, however, that H2O2 is capable of penetrating both chloroplast and
vacuolar membranes, and that anthocyanins can effectively neutralize these ROS in cells
proximate to high light stress (Yamasaki et al. 1996; Gould et al. 2002; Nagata et al. 2003;
Kytridis and Manetas 2006).
There also exists some evidence that anthocyanins function as a solute to decrease
leaf osmotic potential, thereby contributing to osmotic adjustment and maintenance of
turgor pressure in the presence of drought stress during senescence (Chalker-Scott 1999).
In general, anthocyanin synthesis is known to be inducible under conditions which promote
osmotic stress, including high salinity (Dutt et al. 1991 Ramanjulu et al. 1993;
Kaliamoorthy and Rao 1994), drought (Zhi-min et al. 2000; Allen et al. 1999; Spyropoulos
and Mavormmatis 1978), and high sugar treatment (Tholakalabavi et al. 1997, Sakamoto et
al. 1994; Suzuki, 1995 and others). Furthermore, species with high anthocyanin content
7
seem to be common in environments characterized by low soil moisture (Spyropoulos and
Mavormmatis 1978; Schemske and Bierzychudek 2001), and appear to be more tolerant of
drought conditions (Diamantoglou et al. 1989; Knox 1989; Beeson 1992; Paine et al. 1992).
The precise role of anthocyanins within the context of water stress, however, remains
unclear. It has been suggested that anthocyanins may be directly involved in osmotic
regulation, preventing transpirational water loss while helping maintain turgor pressure
(Chalker-Scott 1999, 2002). However, Manetas (2006) reports that, even at very high
concentrations, anthocyanin content in leaf cells is generally much too low to sufficiently
support an osmoregulatory function, and may contribute less than 1% to the osmotic
potential of a leaf. Furthermore, soluble sugars and K+ are ubiquitous among plants while
anthocyanins are not, are less metabolically costly than anthocyanin, and are known to be
the primary contributors to osmotic adjustment in several species (Handa et al. 1983;
Ranney et al. 1991). Another explanation for the association between anthocyanins and
water stress is that anthocyanins are synthesized to alleviate oxidative stress resulting from
low water potentials, either through acting as an antioxidant, or in photoprotection (as
previously described). Under this latter scenario, anthocyanins would be inducible by
water stress, but may not function directly in its alleviation.
Seasonal and ontogenetic occurrence of anthocyanins in photosynthetic tissues
Anthocyanins can be observed in leaves across all ontogenetic stages and seasons
(Figure 1). In most cases, their synthesis corresponds with transient high-light stress, either
due to environmental or ontogenetic limitations to photosynthesis. A decreased capacity
for photosynthesis is known to render plants intrinsically more vulnerable to
8
photoinhibition, due to a reduction in energy sinks available for energy dissipation (i.e.
photochemical quenching, thus increasing their need for photoprotection correspondingly
(Osmond 1981; Powles 1984). For example, immature leaves in the spring and summer
tend to be especially vulnerable to high-light stress due to such factors as: immature
chloroplast structure (Pettigrew and Vaughn 1998; Choinski et al. 2003), reduced capacity
and quantity of photosynthetic enzymes (Miranda et al. 1981a; Pettigrew and Vaughn
1998), and limited stomatal and cellular conductance of CO2 (Miranda et al. 1981b;
Choinski et al. 2003). For this reason, juvenile leaves frequently exhibit biochemical and
structural adaptations that reduce light absorption, including (but not limited to)
anthocyanins (Figure 1A). Seasonal changes may also introduce limitations on
photosynthesis that could render the synthesis of anthocyanin beneficial. Low
temperatures inhibit the biochemical reactions of the Calvin cycle, but do not significantly
affect light capture, which may result in excessive energy captured relative to that used in
carbon fixation. This imbalance leads to an increase in energy and electron transfer to
molecular oxygen, production of radical oxygen species, and increased vulnerability to
photo-oxidative damage (Mittler 2002). This vulnerability to high light under cold
temperatures may have been one of the factors behind evolution of the needle-leaf shape in
conifers during cold, bright glacial periods (Feild et al. 2003), as a cylindrical shape
reduces direct light incidence by the cosine law (Johnson et al. 2005). Similarly,
anthocyanins may represent a more transient photoprotective strategy employed by broadleaved evergreen species, employed only when seasonal changes (e.g. winter) render their
synthesis necessary.
9
Objectives
In the studies presented here, unresolved conflicts or questions associated with a
photoprotective function of anthocyanin pigments are addressed. First, the optical and
photosynthetic effects of anthocyanins in abaxial surfaces of leaves are assessed in two
systems—1) plants which encounter high abaxial irradiance, and 2) plants which grow in
deeply shaded environments characterized by low light—in order to determine whether a
photoprotective function is relevant in either (or both) these cases. Secondly, the dynamic
interaction between anthocyanin synthesis and relative need for photoprotection is
quantified using developing leaves of three deciduous tree species as they transition from
red to green. Finally, the first functional hypothesis behind why some broad-leaf evergreen
species synthesize anthocyanin in winter leaves while others do not, is proposed and tested.
10
Literature Cited
Allen DJ, Nogues S, Morison JIL, Greenslade PD, McLeod AR, and Baker NR. 1999. A
thirty percent increase in UV-B has no impact on photosynthesis in well-watered and
droughted pea plants in the field. Global Change Biology 5: 235-244.
Beeson RC. 1992. Restricting overhead irrigation to dawn limits growth in container-grown
woody ornamentals. HortScience 27: 996-999.
Campbell GS and Norman JM. 1998. Leaf temperature. p.224-229 In: An Introduction to
Environmental Biophysics. Springer, New York, New York.
Chalker-Scott L. 1999. Environmental significance of anthocyanins in plant stress
responses. Photochemistry and Photobiology 70: 1–9.
Chalker-Scott L. 2002. Anthocyanins: Osmoregulators in leaf tissues? Advances in
Botanical Research 37: 103-127.
Choinski JS Jr, Ralph P, and Eamus D. 2003. Changes in photosynthesis during leaf
expansion in Corymbia gummifera. Australian Journal of Botany 51: 111–118.
Cui M, Vogelmann TC, and Smith WK. 1991. Chlorophyll and light gradients in sun and
shade leaves of Spinacia oleracea. Plant, Cell & Environment 14: 493-500.
Davies KM and Schwinn KE. 2003. Transcriptional regulation of secondary metabolism.
Functional Plant Biology 30: 913–925.
Diamantoglou S, Rhizopoulou S, and Kull U. 1989. Energy content, storage substances,
and construction and maintenance costs of Mediterranean deciduous leaves. Oecologia
81: 528-533.
Dutt SK, Bal AR, and Bandyopadhyay AK. 1991. Salinity induced chemical changes in
Casuarina equisetifolia Forst. Egyptian Journal of Soil Science 31: 57-63.
Feild TS, Lee DW, and Holbrook NM. 2001. Why leaves turn red in autumn. The role of
anthocyanins in senescing leaves of red-osier dogwood. Plant Physiology 127: 566-574.
Feild TS, Arens NC, and Dawson TE. 2003. The ancestral ecology of angiosperms:
emerging perspectives from extant basal lineages. In D.A. Ackerly and R. C. Monson [eds.],
Evolution of functional traits in plants. International Journal of Plant Sciences 164
(Supplement): S129–S142.
Gould KS, Kuhn DN, Lee DW, and Oberbauer SF. 1995. Why leaves are sometimes red.
Nature 378: 241-242.
11
Gould KS, McKelvie J, and Markham KR. 2002. Do anthocyanins function as antioxidants
in leaves? Imaging of H2O2 in red and green leaves after mechanical injury. Plant, Cell &
Environment 25: 1261-1269.
Gould KS. 2004. Nature’s Swiss Army Knife: The diverse protective roles of anthocyanins
in leaves. Journal of Biomedicine and Biotechnology 5: 314-320.
Handa S, Bressan RA, Handa AK, Carpita NC, and Hasegawa PM. 1983. Solutes
contributing to osmotic adjustment in cultured plant cells adapted to water stress. Plant
Physiology 73: 834-843.
Holton TA and Cornish EC. 1995. Genetics and biochemistry of anthocyanin biosynthesis.
The Plant Cell 7: 1071-1083.
Hughes NM, Neufeld HS, and Burkey KO. 2005. Functional role of anthocyanins in highlight winter leaves of the evergreen herb Galax urceolata. New Phytologist 168: 575-587.
Hughes NM and Smith WK. 2007. Seasonal photosynthesis and anthocyanin production in
ten broadleaf evergreen species. Functional Plant Biology 34: 1072-1079.
Iwashina T. 2000. The structure and distribution of the flavonoids in plants. Journal of
Plant Research 113: 287-299.
Johnson DM, Smith WK, Vogelmann TC, and Brodersen CR. 2005. Leaf architecture
and direction of incident light influence mesophyll fluorescence profiles. American
Journal of Botany 92: 1425-1431.
Kaliamoorthy S and Rao AS. 1994. Effect of salinity on anthocyanin accumulation in the
root of maize. Indian Journal of Plant Physiology 37: 169-170.
Knox GW. 1989. Water use and average growth index of five species of container grown
woody landscape plants. Journal of Environmental Horticulture 7: 136-139.
Kyparissis I, Grammatikopoulos G, and Manetas Y. 2007. Leaf morphological and
physiological adjustments to the spectrally selective shade imposed by anthocyanins in
Prunus cerasifera. Tree Physiology 27: 849-857.
Kytridis V-P and Manetas Y. 2006. Mesophyll versus epidermal anthocyanins as
potential in vivo antioxidants: evidence linking the putative antioxidant role to the
proximity of the oxy-radical source. Journal of Experimental Botany 57: 2203-2210
Lee DW, Lowry JB, and Stone BC. 1979. Abaxial anthocyanin layer in leaves of tropical
rain forest plants: enhancer of light capture in deep shade. Biotropica 11: 70-77.
Lee DW, Brammeier W, and Smith AP. 1987. The selective advantages of anthocyanins in
developing leaves of mango and cacao. Biotropica 19: 40-49.
12
Lee, DW, O’Keefe J, Holbrook NM, and Field TS. 2003. Pigment dynamics and autumn
leaf senescence in a New England deciduous forest, eastern USA. Ecological Research 18:
677-694.
Lee DW and Gould KS. 2002. Anthocyanins in leaves and other vegetative organs: An
introduction. Advances in Botanical Research 37: 2-16.
Mancinelli, AL. 1993. The photoregulation of anthocyanin synthesis. In: Shropshire Jr. W
and Mohr H (eds.), Encyclopedia of Plant Physiology. New Series, Vol. 16B. Springer,
Berlin. Pp. 640-661.
Manetas Y, Petropoulou Y, Psaras GK, and Drinia A. 2003. Exposed red (anthocyanic)
leaves of Quercus coccifera display shade characteristics. Functional Plant Biology 30:
265-270.
Manetas Y. 2006. Why some leaves are anthocyanic and why most anthocyanic leaves are
red. Flora 201: 163–177.
Marrs KA, Alfenito MR, Lloyd AM, and Walbot V. 1995. A glutathione S-transferase
involved in vacuolar transfer encoded by the maize gene Bronze-2. Nature 375: 397–400.
Merzlyak MN and Chivkunova OB. 2000. Light-stress-induced pigment changes and
evidence for anthocyanin photoprotection in apples. Journal of Photochemistry and
Photobiology 55: 155-163.
Miranda V, Baker NR, and Long SP. 1981a. Limitation of photosynthesis in different
regions of the Zea mays leaf. New Phytologist 89: 179–190.
Miranda V, Baker NR, and Long SP. 1981b. Anatomical variation along the length of the
Zea mays leaf in relation to photosynthesis. New Phytologist 88: 595–605.
Mittler R. 2002. Oxidative stress, antioxidants, and stress tolerance. Trends in Plant
Science 7: 405-410.
Nagata T, Todoriki S, Masumizu T, Suda I, Furuta S, Du Z, and Kikuchi S. 2003. Levels of
active oxygen species are controlled by ascorbic acid and anthocyanin in Arabidopsis.
Journal of Agriculutural Food Chemistry 51: 2992-2999.
OsmondCB. 1981. Photorespiration and photoinhibition: some implications for the
energetics of photosynthesis. Biochimica et Biophysica Acta 639: 77–98.
Ougham HJ, Thomas H, and Archetti M. 2008. Origin and evolution of autumn colours.
New Phytologist 179: 9-13.
13
Paine TD, Hanlon CC, Pittenger DR, Ferrin DM, and Malinoski MK. 1992. Consequences
of water and nitrogen management on growth and aesthetic quality of drought-tolerant
woody landscape plants. Journal of Environmental Horticulture 10: 94-99.
Pettigrew WT and Vaughn KC. 1998. Physiological, structural, and immunological
characterization of leaf and chloroplast development in cotton. Protoplasma 202: 23–37.
Pietrini F, Iannelli MA, and Massacci A. 2002. Anthocyanin accumulation in the
illuminated surface of maize leaves enhances protection from photo-inhibitory risks at low
temperature, without further limitation to photosynthesis. Plant, Cell & Environment 25:
1251–1259.
Powles SB.1984. Photoinhibition of photosynthesis induced by visible light. Annual
Review of Plant Physiology 35: 15–44.
Ramanjulu S, Veeranjaneyulu K, and Sudhakar C. 1993. Physiological changes induced by
NaCl in mulberry var. Mysore local. Indian Journal of Plant Physiology 36: 273-275.
Rolland F, Moore B, and Sheen J. 2002. Sugar sensing and signaling in plants. Plant Cell
(Suppl) 14: S185–S205.
Rook F and Bevan MW. 2003. Genetic approaches to understanding sugar response
pathways. Journal of Experimental Botany 54: 495–501.
Sakamoto K, Kumiko I, Sawamura K, Kyoko H, Yoshihisa A, Takafumi Y, and Tsutomu F.
1994. Anthocyanin production in cultured cells of Aralia cordata Thumb. Plant, Cell
Tissue, and Organ Culture 36: 21-26.
Schemske DW and Bierzychudek P. 2001. Evolution of flower color in the desert annual
Linanthus parryae: Wright revisited. Evolution 55: 1269-1282.
Smeekens S. 2000. Sugar-induced signal transduction in plants. Annual Review of Plant
Physiology and Plant Molecular Biology 51: 49–81.
Smillie RM and Hetherington SE. 1999. Photoabatement by anthocyanin shields
photosynthetic systems from light stress. Photosynthetica 36: 451-463.
Smith AM. 1909. On the internal temperature of leaves in tropical insolation, with special
reference to the effects of their colour on temperature; also observations on the periodicity
of the appearance of young coloured leaves of trees growing in Peradeniya
Gardens. Annals of the Royal Botanical Gardens, Peradeniya 5: 229–298.
Spyropoulos CG and Mavrommatis M. 1978. Effect of water stress on pigment formation
in Quercus species. Journal of Experimental Botany 29: 473-477.
Sturgeon KB and Mitton JB. 1980. Cone color polymorphism associated with elevation
14
in white fir, Abies concolor, in southern Colorado. American Journal of Botany 67: 1040
1045.
Suzuki M. 1995. Enhancement of anthocyanin accumulationby high osmotic stress and
low pH in grape cells (Vitis hybrids). Journal of Plant Physiology 147: 152-155.
Teng S, Keurenties J, Bentsink L, Koornneef M, and Smeekens S. 2005. Sucrose-specific
induction of anthocyanin biosynthesis in Arabidopsis requires the MYB75/PAP1 gene.
Plant Physiology 139: 1840-1852.
Tholakalabavi A, Zwiazek JJ, and Thorpe TA. 1997. Osmotically-stressed poplar cell
cultures: anthocyanin accumulation, deaminase activity, and solute composition. Journal of
Plant Physiology 151: 489-496.
Vanderauwera S, Zimmermann P, Rombauts S, Vandenabeele S, Langebartels C, Gruissem
W, Inze D, and Van Breusegem F. 2005. Genome-wide analysis of hydrogen peroxideregulated gene expression in Arabidopsis reveals a high light-induced transcriptional
cluster involved in anthocyanin biosynthesis. Plant Physiology 139: 806–821.
Weiss D. 2000. Regulation of flower pigmentation and growth: multiple signaling
pathways control anthocyanin synthesis in expanding petals. Physiologia Plantarum 110:
152–157.
Yamasaki H, Uefuji H, and Sakihama Y. 1996. Bleaching of the red anthocyanin induced
by superoxide radical. Archives of Biochemistry and Biophysics 332: 183-186.
Zhi-min, Y, Shao-jian Z, Ai-tong H, You-fei Z, and Jing-yi Y. 2000. Response of
cucumber plants to increased UV-B radiation under water stress. Journal of Environmental
Science 12: 236-240.
15
Figure 1.1
Anthocyanins may be observed in leaves during all life stages and seasons. Examples: (A)
Quercus alba synthesizes anthocyanin in juvenile leaves during spring; (B) Saxifraga
michauxii exhibits abaxial reddening during summer months in mature leaves; (C) Acer
saccharum synthesizes anthocyanin in senescing leaves during autumn; (D) the evergreen
herb Galax urceolata exhibits reddening due to anthocyanin accumulation in mature leaves
during winter.
16
Figure 1.1
17
Figure1.2
The biosynthetic pathway for anthocyanin, beginning at malonyl-CoA and p-coumaroylCoA (the precursors of all flavonoids), showing how the three general colors of
anthocyanin are synthesized (from Holton and Cornish, 1995).
18
Figure 1.2
19
Figure 1.3. (A) Photoinduction of anthocyanin synthesis in Lonicera japonica, (B) the
direct relationship between light intensity and anthocyanin accumulation, illustrated in
Galax urceolata.
20
Figure 1.3
21
Figure 1.4. (A) Vacuolar sequestration of anthocyanin in palisade mesophyll cells of
Rhododendron spp. (B) Adaxial anthocyanin in palisade mesophyll cells of Leucothoe
fontanesiana.
22
Figure 1.4
23
CHAPTER II
ATTENUATION OF INCIDENT LIGHT IN GALAX URCEOLATA (DIAPENSIACEAE):
CONCERTED INFLUENCE OF ADAXIAL AND ABAXIAL ANTHOCYANIC
LAYERS ON PHOTOPROTECTION
Nicole M. Hughes and William K. Smith
The following manuscript was published in American Journal of Botany, volume 94, pages
784-790, 2007, and is reprinted with permission.
Stylistic variations are due to the
requirements of the journal. N.M. Hughes performed the field work, data analysis, and
prepared the manuscript. W.K. Smith provided logistical support, advice and ideas, and
editorial assistance.
24
Abstract
Although anthocyanin coloration in lower (abaxial) leaf cells has been documented
for numerous species, the functional significance of this character has not been
comprehensively investigated according to habitat or leaf orientation. In the current study,
we demonstrate that abaxial anthocyanin may function as a photoprotectant, similarly to its
purported role in upper (adaxial) cells, in leaves vulnerable to high irradiance incident on
abaxial surfaces. Spectral scans were derived for Galax urceolata leaves exhibiting the
following phenotypes: abaxial or adaxial anthocyanin only, abaxial and adaxial
anthocyanin, and no anthocyanin. To determine whether anthocyanins conferred protection
from photoinhibition, maximum photosystem II efficiencies of red (anthocyanic) and green
(acyanic) surfaces were compared during and after exposure to photoinhibitory conditions.
Leaves were either positioned with their adaxial surfaces facing the light source or inverted
to expose abaxial surfaces. Spectral scans showed increased absorptance of 500-600 nm
wavelengths by red surfaces (consistent with the absorbance spectrum of anthocyanin),
regardless of whether that surface was abaxial or adaxial. Leaves with anthocyanin in
either illuminated surface were also photoinhibited less than leaves lacking anthocyanin in
that surface. These results suggest that anthocyanic layers reduce absorbed sunlight in the
mesophyll not only for adaxial surfaces, but abaxial as well. Adaxial/abaxial anthocyanin
plasticity may therefore be adaptive in high-light environments or during light-sensitive
developmental stages where leaf orientation and/or substrate albedo are variable.
25
Introduction
Anthocyanins have received considerable attention in the recent literature due to
their proposed role as light-attenuators in leaves vulnerable to high light stress (DrummHerrel and Mohr, 1985; Landry et al., 1995; Smillie and Hetherington, 1999; Gould et al.,
2000; Feild et al., 2001; Gould et al., 2002; Pietrini et al., 2002; Hoch et al., 2003; Neill
and Gould, 2003; Manetas et al., 2003, Hughes et al., 2005; Karageorgou and Manetas,
2006). By absorbing blue-green light that may otherwise be absorbed by chlorophyll b in
subjacent mesophyll cells, anthocyanins are thought to relieve the potential damage to the
photosystems that occurs when leaves are exposed to more light energy than can be
effectively assimilated for photosynthesis or dissipated as heat. The physiological
advantage that this confers has been evidenced in studies comparing light capture
efficiency of anthocyanic and cyanic leaves under photoinhibitory conditions. When red
and green leaf surfaces (typically adaxial) are exposed to high light stress, red leaves tend
to be less photoinhibited than green leaves when other photoprotective mechanisms are
equal (Feild et al., 2001; Hoch et al., 2003; Hughes et al., 2005). In the current study, we
test whether abaxial anthocyanin may perform the same function in leaves where the
abaxial surface is incident to high light rather than (or in addition to) the adaxial surface.
Exposure to high abaxial irradiance occurs under a broad range of conditions: in
environments with high substrate albedos (e.g. snow and sand), among plants with inclined
leaf angles that expose abaxial surfaces to ambient sunlight, and in species where
developing juvenile leaves emerge with abaxial surfaces facing outwards (Drumm-Herrel
and Mohr, 1985; Sherwin and Farrant, 1998; personal observation). However, little is
known about adaptations of abaxial cells to withstand high irradiance, which is surprising
26
given the increased sensitivity of abaxial surfaces relative to adaxial in dorsiventrally
asymmetric leaves (Sun et al., 1996; Sun and Nishio, 2001). Although the synthesis of
anthocyanin in abaxial cells in response to high light stress has been reported (Barker et al.,
1997; Sherwin and Farrant, 1998; Pietrini et al., 2002; Hughes et al., 2005), and
development of photostability has been shown to occur more quickly in seedlings with
abaxial anthocyanin (Drumm-Herrel and Mohr, 1985), no studies have directly tested the
prospect that these pigments decrease susceptibility to photoinhibition through light
attenuation in abaxial cells.
Previous studies on abaxial anthocyanin have focused almost exclusively on
understory species adapted to low light environments, where red and purple abaxial
surfaces are also a common morphological character (Richards 1952; Lee et al., 1979; Lee
and Graham, 1986; Lee and Collins, 2001). Under these conditions it has been suggested
that abaxial anthocyanin functions to back-scatter red wavelengths that enter the leaf from
the adaxial surface, thereby increasing light capture efficiency in low-intensity and poorlight quality environments (Lee et al., 1979). However, this interpretation has not been
well-supported, and seems insufficient to explain the appearance of anthocyanin in abaxial
cells of leaves experiencing high light stress (where an increase in light capture would
likely exacerbate photoinhibition). Under such conditions, it is more likely that
anthocyanins are functioning as light attenuators to reduce photoinhibition of abaxial cells.
To determine whether abaxial anthocyanin functions to decrease susceptibility to
photoinhibition similarly to adaxial anthocyanin, we take advantage of the phenotypic
plasticity found in the evergreen understory herb Galax urceolata (Poir.) Brummitt for
synthesizing anthocyanin on either or both leaf surfaces receiving moderately high levels of
27
irradiance (i.e. > 750 µmol · m-2 · s-1) during the winter. In its natural environment, G.
urceolata leaves are typically oriented with adaxial surfaces facing canopy gaps, with leaf
angles ranging from 0° to 90° (most commonly between 40° and 60°). Although highest
irradiances are typically incident on adaxial surfaces (resulting in adaxial anthocyanin),
abaxial surfaces of leaves with more vertical angles often experience relatively high
irradiance from snow albedo and direct and diffuse light, resulting in anthocyanin synthesis
in abaxial surfaces as well. Less frequently, leaf litter or debris will cause leaves to be
entirely inverted, and only abaxial surfaces will appear red. By comparing the optical
properties and ecophysiology of leaves with and without adaxial and abaxial anthocyanin,
we demonstrate that anthocyanin located within abaxial tissues attenuates light in the same
manner as anthocyanin located within adaxial cells, effectively reducing photoinhibition of
abaxial leaf tissues vulnerable to high light stress.
28
Materials and Methods
To compare optical profiles of abaxially- and adaxially-anthocyanic leaves, spectral
scans were derived for leaves with the following pigment patterns: entirely green (green
adaxial and green abaxial), bicolored (either green adaxial with red abaxial, or red adaxial
with green abaxial), or entirely red (red adaxial, red abaxial) (Fig. 1). Three leaves of each
type were removed from the field and stored under cold, moist conditions until analysis
later that day. Spectral scans for reflectance and transmittance of photosynthetically active
radiation (PAR), representing 400-700 nm, of both adaxial and abaxial surfaces were
derived using a Li-Cor 1800 spectroradiometer with external integrating sphere (Li-Cor,
Inc., Lincoln, Nebraska, USA). Measurements were taken at 1-nm intervals. Absorptance
was calculated as (1 – reflectance – transmittance). The area tested was marked, and leaf
thickness and pigment concentrations were subsequently quantified. Leaves were all first
year leaves obtained from a wooded area in Jonas Ridge, North Carolina, USA (35°57’20”
N, 81°53’55” W) in February 2006.
Leaf thickness was measured using digital calipers with 0.01 mm precision.
Chlorophylls were extracted by placing three 0.33 cm2 leaf discs in 3 mL N,N’ dimethylformamide in the dark for 24 h. Concentrations were determined using the
equations described by Porra (2002). Anthocyanins were extracted by placing three discs
in 3 mL 6M HCl:H2O:MeOH (7:23:70) in the dark at 4oC for 24 h. Following extraction, 1
mL of the solution was removed, and to it 1 mL water and 1mL of chloroform were added
to separate anthocyanins (insoluble in chloroform) from chlorophylls. The solutions were
centrifuged for 15 minutes at 2630 g and 1 mL of the top layer (containing anthocyanins)
was used for analysis. Anthocyanin concentration was determined as A530, using an
29
extinction coefficient of 30 000 l mol-1 cm-1 (Murray and Hackett, 1991). Prior to
extraction, all leaf discs were submerged in liquid nitrogen to infiltrate cuticular waxes and
disrupt cell membranes.
Chlorophyll a and b concentrations, ratios, total chlorophyll content, and leaf
thickness of the four leaf groups were compared using a single factor ANOVA with TukeyKramer test for means difference, using JMP statistical software (SAS Institute, Cary, NC)
(Zar, 1999). Anthocyanin concentrations were compared using a single factor ANOVA
with planned contrast. Completely green leaves were predicted a priori to exhibit
significantly less anthocyanin than all other groups, and completely red leaves were
predicted to exhibit significantly more.
For microscopic imaging, leaves were sectioned into 50-100 µm sections using a
vibratome. Sections were mounted on a Zeiss Axioplan upright microscope (Carl Zeiss
Inc., Thornwood, NY), viewed using under differential interference contrast (DIC)
microscopy, and images were captured using a Hamamatsu C5810 three-chip cooled color
CCD camera (Hamamatsu Photonics; Hamamatsu City, Japan). Photographs of leaf
sections were rotated and adjusted for brightness and image sharpness using Adobe
Photoshop CS (Adobe Systems, San Jose, California).
Another set of G. urceolata leaves was used to compare maximum light capture
efficiency (Fv/Fm) of the leaf types during and following exposure to photoinhibitory
conditions. Leaves were derived from the same plots described above, also in February.
Fluorescence measurements were taken using a Handy-PEA 1000 fluorescence analyzer
(Hansatech Institute, Cambridge, UK), emitting a two second, 3 mmol · m-2 · s-1 saturating
pulse. Prior to measurement, plants were dark-adapted for 30 min using Handy-PEA leaf
30
clips. Five leaves with red adaxial surfaces and five leaves with green adaxial surfaces
(both having green abaxial surfaces) were used to test adaxial response to high light stress.
Abaxial surfaces were tested in a separate experiment using five leaves for each of three
phenotypes: green adaxial, green abaxial; red adaxial, red abaxial; red adaxial, green
abaxial. Experiments differed only in the surface of the leaf exposed to the light source.
Because results from abaxial experiments showed no effect of anthocyanin in the nonilluminated surface on photoprotection, it did not seem necessary to test all four
combinations of anthocyanic layers in this study.
Leaves were collected from the field, and petioles were re-cut underwater and
remained submerged until experimentation. To equalize starting Fv/Fm values, all leaves
were placed in low stress environments for four days until no significant difference
between Fv/Fm values of leaf groups was observed (for detailed description of why field
Fv/Fm values of red and green leaves differ and why standardization of Fv/Fm is necessary,
see Hughes et al., 2005). During this time, leaves with anthocyanic surfaces were kept at
constant temperature (18°C) and exposed to 10 h of low light (175 μmol · m-2 · s-1),
provided by a 1000 W metal halide bulb equipped with water bath and neutral density
shade cloth to obtain desired PPFDs, to allow Fv/Fm to recover to values similar to those of
green leaves. Green leaves were placed under similar light conditions, but exposed to field
temperatures (−10 to 15°C) to prevent Fv/Fm values from exceeding 0.6-0.7 (which red
leaves could not recover to during the winter). Once values of Fv/Fm were no longer
significantly different between the leaf groups, all leaves were placed in the outdoor,
protected enclosure overnight. At 7 am the following morning, leaves were exposed to a
high stress treatment, consisting of high light (1150 µmol · m-2 · s-1 provided by a 1000 W
31
metal halide bulb) and low temperatures (circulating outside air, with daily temperatures
ranging from 0 to 10oC during the day, and −15 to 0oC during the night). High light stress
was applied for 10 h per day over 3 consecutive days. On the fourth day, leaves were
transferred into a low stress environment (same conditions as those described above to pretreat red leaves) until Fv/Fm recovered to near starting values. Fluorescence readings were
taken at 5 pm each day throughout the experiment. A repeated measures MANOVA was
used to compare fluorescence between leaf types during the high light treatments.
32
Results
Levels of chlorophyll a and b, total chlorophyll content, chlorophyll a:b, and leaf
thickness did not significantly differ between the four leaf phenotypes in G. urceolata (p=
0.96, 0.85, 0.85, 0.27, and 0.26 respectively) (Table 1). Leaves with anthocyanin occurring
on both adaxial and abaxial surfaces had significantly higher anthocyanin content than the
other 3 leaf types (p< 0.01), exhibiting 2-fold higher anthocyanin concentration than
bicolored leaves, and >7-fold higher concentration than entirely green leaves (which
contained virtually no anthocyanin). Adaxially and abaxially-bicolored leaves did not
significantly differ in anthocyanin concentration (p= 0.86), but did contain significantly
more anthocyanin than green leaves (p= 0.037) (Table 1).
Leaves with anthocyanin in both surfaces absorbed the most blue-green to yellow
(500-600 nm) light of the four leaf types, absorbing up to 20% more of the green
wavelengths than entirely green leaves, and 2-3% more than bicolored leaves (Figs. 2A and
B). Absorption spectra of red and green bicolored leaves were similar to those of entirely
red leaves when light was shone on the anthocyanic surface, though absorption was
reduced by roughly 10% when the acyanic surface was illuminated. However, absorptance
of green wavelengths was still much higher (by up to 10%) than entirely green leaves. The
increased absorptance of green light by anthocyanic tissues resulted in a dramatic decrease
in transmittance of green wavelengths for anthocyanic leaves relative to entirely green
leaves (Figs. 2C and D), with entirely red leaves reducing transmittance of blue-green light
(500-550 nm) to 0%, and bicolored leaves to 2-3% (10% lower than green leaves). The
increased absorptance of green light by anthocyanin also corresponded to a decrease in
reflectance of these wavelengths when the red surface was illuminated (by up to 10%),
33
though reflectance was not affected when light was incident on the green surface (Figs. 2E
and F). In all leaves containing anthocyanin, roughly 1% more incident red light (600-675
nm) was reflected from the leaf surface compared to entirely green leaves, regardless of
which surface was illuminated (acyanic or anthocyanic).
A comparison of spectral scans of abaxial and adaxial surfaces showed that leaves
absorbed more light (5-7% between 450 and 600 nm wavelengths) when light was incident
upon the adaxial surface, irregardless of pigmentation (Figs. 2A and B). Furthermore,
leaves reflected more light of all wavelengths when the light was incident on the abaxial
surface (Figs. 2D and E). However, transmittance across the leaf was not affected by the
directionality of the light source (Figs. 2C and D).
Green leaf Fv/Fm declined by 86% compared to only 55% for red leaves following
exposure to photoinhibitory conditions (Fig. 3A), a difference which was highly significant
(p<0.0001). This trend was also observed for abaxial surfaces (Fig. 3B, p<0.001), where
red undersides exhibited significantly less of a decline in Fv/Fm than green undersides
(regardless of whether the opposite surface was red or green). Photoinhibition of red
abaxial cells was approximately 10% higher than that of red adaxial cells, though patterns
of photoinhibition between all green surfaces (adaxial and abaxial) were similar. Recovery
from photoinhibition also appeared to be affected by anthocyanin, as Fv/Fm values of red
adaxial surfaces recovered to near-starting values after 1 d, while green adaxial surfaces
required 5 d. This trend was also observed with red abaxial surfaces compared to green.
34
Discussion
Analyses of leaf pigments and thicknesses (Table 1) showed that the leaf
phenotypes measured here did not significantly differ in chlorophyll content or leaf
thickness, only anthocyanin concentration. Therefore, any deviations in optical properties
between leaf types were most likely due to the effect of anthocyanin.
Consistent with known spectral characteristics of anthocyanin, spectral scans
indicated that anthocyanins affected leaf optical properties most apparently through their
attenuation of blue-green to yellow (500-600 nm) light (Fig. 2), with peak absorptance
occurring between 530-550 nm. When light was incident on red surfaces, absorptance of
these wavelengths increased by up to 20%, regardless of whether that surface was abaxial
or adaxial, or whether the opposite surface contained anthocyanin or not. This suggests
that abaxial and adaxial anthocyanic layers may function independently to protect adjacent
mesophyll cells, or in concert to protect the leaf entire. Increased absorptance of bluegreen to yellow light also corresponded with decreased transmittance (Figs. 2C and D) and
reflectance (Figs. 2E and F) of these wavelengths. Because absorptance of blue-green light
by anthocyanin has been shown to protect leaves from photoinhibition (Feild et al., 2001;
Hughes et al., 2005), these data support the idea that anthocyanins on either leaf surface
confer photoprotection.
Because leaves with high anthocyanin concentrations appear red, the
photoprotective effect of anthocyanin may be misattributed to an increase in reflectance of
red light. Because red light is preferentially absorbed by chlorophyll, increasing
reflectance of these wavelengths would indeed be effective in curtailing photoinhibition.
However, as show in Figures 2E and F, the presence of anthocyanin does not necessarily
35
increase the percentage of red light reflected by the leaf. Instead, leaves appear red due to
the diminished reflectance of other wavelengths (primarily green and blue). As a result, the
reflectance peak shifts from green to red as leaves accumulate anthocyanin, though the
magnitude of reflectance of red wavelengths does not markedly change (i.e. greater than
1%). In addition, previous work found no difference in photoinhibition between red and
green G. urceolata leaves illuminated with red light (Hughes et al., 2005). Therefore, any
apparent increase in reflectance of red wavelengths afforded by anthocyanin does not
convey a photoprotective effect in this species.
When the green surfaces of bicolored leaves were illuminated, it was found that
anthocyanins were still able to intercept up to 10% of green wavelengths that would
otherwise have been transmitted through the leaf (Fig. 2A and B). Since an increase in
sunlight absorptance often translates into an increase in leaf temperature, it has been
proposed in previous studies that abaxial pigment layers may function to elevate leaf
temperature (Smith, 1909; Lee et al., 1979; Lee et al., 1987). Although this function would
seem adaptive in plants susceptible to low temperature photoinhibition, several studies of
abaxially anthocyanic plants (including G. urceolata) have found no difference in leaf
temperature between anthocyanic and acyanic leaves (Lee et al., 1979; Lee et al., 1987;
Hughes, 2004).
The effects of anthocyanin on maximum photosystem II efficiency (Fv/Fm) during
photoinhibitory conditions are shown in Figure 3. Leaves with red surfaces appeared less
susceptible to low temperature photoinhibition (as evidenced by higher Fv/Fm) during the
high stress treatment than leaves with green surfaces, regardless of whether that surface
was adaxial (3A) or abaxial (3B). Furthermore, anthocyanic surfaces recovered to starting
36
Fv/Fm values more quickly than acyanic surfaces, suggesting a lesser degree of
photodamage had been incurred. In a previous study on G. urceolata by the authors
(Hughes et al., 2005), it was further shown that photoinhibition of red and green adaxial
surfaces were similar under red light (which anthocyanins do not absorb), but under green
light (which anthocyanins absorb strongly) red surfaces were significantly less
photoinhibited than green. Therefore, we can presume in the case of abaxial anthocyanins
as well, that any difference in Fv/Fm is due to the presence of anthocyanin, and not, for
example, a difference in the pre-treatments (which would have been evident under both red
and green light conditions). Finally, it was also observed that the presence of anthocyanin
in the non-illuminated surface did not appear to have an effect on Fv/Fm (3B), again
suggesting an independent function of these layers on photoprotection of their respective
surface.
An additional trend that is apparent from these data is the increased susceptibility of
red abaxial cells to photoinhibition compared to red adaxial cells. This is most likely due to
lower anthocyanin content in abaxial relative to adaxial cells of leaves tested in this
experiment (data not shown), as natural leaf angles of this species generally cause adaxial
surfaces to receive the greatest incoming radiation. This is most likely due to lower
anthocyanin content in abaxial relative to adaxial cells (data not shown) due to less intense
incident PAR (natural leaf angles of this species cause adaxial surfaces receive the greatest
incoming radiation). Another trend worth noting is that the protection afforded by
anthocyanin appears to be independent of the presence of anthocyanin on the opposite
surface (3B), again suggesting that leaf surfaces function independently with regards to
protecting tissues from light incident on that surface.
37
Given the above findings, which support the idea that abaxial anthocyanins function
in light attenuation, what might be the function of abaxial anthocyanins in leaves of deeply
shaded understory plants (where abaxial red coloration is most common)? As mentioned
previously, although no studies have been able to provide a well-supported explanation, it
is possible that abaxial anthocyanins are acting as light-attenuators in shade plants as well.
For example, it is possible that abaxial anthocyanin protects the mesophyll from high
ambient light (either diffuse or caused by sunflecks) striking the extremely-dark adapted
abaxial surfaces. Another possibility, based on current research by the authors on Begonia
spp., is that abaxial anthocyanins attenuate internally-scattered green light (entering the leaf
from the adaxial surface), thereby buffering internal light levels during high-intensity
sunflecks. This function is consistent with the findings of Gould et al. (1995), where
abaxially-red tropical understory plants had significantly higher Fv/Fm than abaxially-green
conspecifics.
In conclusion, the results reported here for Galax urceolata support the idea that
anthocyanins located in either adaxial or abaxial surfaces may confer a photoprotective
advantage in tissues exposed to photoinhibitory conditions by effectively intercepting PAR
incident on the respective surface. Moreover, the abaxial/adaxial anthocyanic plasticity
observed in G. urceolata and numerous other species may be an important adaptation in
photoinhibitory environments and/or during light-sensitive developmental stages where
leaf orientation and/or substrate albedo are variable.
38
Literature Cited
BARKER D. H., G. G. R. SEATON, AND S. A. ROBINSON. 1997. Internal and external
photoprotection in developing leaves of the CAM plant Cotyledon orbiculata. Plant, Cell
and Environment 20: 617-624.
DRUMM-HERREL H., AND H. MOHR. 1985. Photosensitivity of seedlings differing in
their potential to synthesize anthocyanin. Physiologia Plantarum 64: 60-66.
FEILD, T. S., D. W. LEE, AND N. M. HOLBROOK. 2001. Why leaves turn red in autumn.
The role of anthocyanins in senescing leaves of red-osier dogwood. Plant Physiology 127:
566–574.
GOULD K.S., D.N. KUHN, D.W. LEE, AND S.F. OBERBAUER. 1995. Why leaves are
sometimes red. Nature 378: 241 – 242.
GOULD, K. S., K. R. MARKHAM, R. H. SMITH, AND J. J. GORIS. 2000. Functional
role of anthocyanins in the leaves of Quitinia serrata A. Cunn. Journal of Experimental
Botany 51: 1107–1115.
GOULD, K. S., T. C. VOGELMANN, T. HAN, AND M. J. CLEARWATER. 2002.
Profiles of photosynthesis within red and green leaves of Quintinia serrata. Physiologia
Plantarum 116: 127-133.
HOCH W. A., E. L. SINGSAAS, AND B. H. MCCOWN. 2003. Resorption protection.
Anthocyanins facilitate nutrient recovery in autumn by shielding leaves from potentially
damaging light levels. Plant Physiology 133: 1296-1305.
HUGHES N. M. 2004. Functional role of anthocyanins in high light winter leaves of the
evergreen herb Galax urceolata. MSc thesis, Boone, NC, USA: Appalachian State
University.
HUGHES N. M., H. S. NEUFELD, AND K. O. BURKEY. 2005. Functional role of
anthocyanins in high-light winter leaves of the evergreen herb Galax urceolata. New
Phytologist 168: 575-587.
KARAGEORGOU P., AND Y. MANETAS. 2006. The importance of being red when
young: anthocyanins and the protection of young leaves of Quercus coccifera from insect
herbivory and excess light. Tree Physiology 26: 613-621.
LANDRY L. G., C. C. S. CHAPPLE, AND R. L. LAST. 1995. Arabidopsis mutants
lacking phenolic sunscreens exhibit enhanced ultraviolet-B injury and oxidative damage.
Plant Physiology 109: 1159–1166.
39
LEE D. W., J. B. LOWRY, AND B. C. STONE. 1979. Abaxial anthocyanin layer in leaves
of tropical rain forest plants: enhancer of light capture in deep shade. Biotropica 11(1): 7077.
LEE D. W., AND R. GRAHAM. 1986. Leaf optical properties of rainforest sun and
extreme shade plants. American Journal of Botany 73: 1100-1108.
LEE D. W., S. BRAMMEIER, AND SMITH A.P. 1987. The selective advantages of
anthocyanins in developing leaves of mango and cacao. Biotropica 19: 40-49.
LEE D. W., AND T. M. COLLINS. 2001. Phylogenetic and ontogenetic influences on the
distribution of anthocyanins and betacyanins in leaves of tropical plants. International
Jounral of Plant Sciences 162: 1141-1153.
MANETAS Y., Y. PETROPOULOU, G. K. PSARAS, AND A. DRINIA. 2003. Exposed
red (anthocyanic) leaves of Quercus coccifera display shade characteristics. Functional
Plant Biology 30:265-270.
MURRAY J. R., AND W. P. HACKETT. 1991. Dihydroflavonol reductase activity in
relation to differential anthocyanin accumulation in juvenile and mature phase Hedera helix
L. Plant Physiology 97: 343-351.
NEILL S. O., AND K. S. GOULD. 2003. Anthocyanins in leaves: light attenuators or
antioxidants? Functional Plant Biology 30: 865–873.
PIETRINI F., M. A. IANNELLI, AND A. MASSACCI. 2002. Anthocyanin accumulation
in the illuminated surface of maize leaves enhances protection from photo-inhibitory risks
at low temperature, without further limitation to photosynthesis. Plant, Cell & Environment
25: 1251–1259.
PORRA R. J. 2002. The chequered history of the development and use of simultaneous
equations for the accurate determination of chlorophylls a and b. Photosynthesis Research
73: 149-156.
RICHARDS P.W. 1952. The tropical rainforest. Cambridge University Press, London.
SHERWIN H. W., AND J. M. FARRANT. 1998. Protection mechanisms against excess
light in the resurrection plants Craterostigma wilmsii and Xerophyta viscosa. Plant Growth
Regulation 24: 203-210.
SMILLIE R. M., AND S. E. HETHERINGTON. 1999. Photoabatement by anthocyanin
shields photosynthetic systems from light stress. Photosynthetica 36: 451–463.
SMITH A. M. 1909. On the internal temperatures of leaves in tropical insolation, with
special reference to the effect of their colour on temperature; also observations on the
40
periodicity of the appearance of young coloured leaves of trees growing in Peradinaya
Gardens. Annals of the Royal Botanical Garden. Peradinaya 5: 229-297.
SUN J., J. N. NISHIO, AND T. C. VOGELMANN. 1996. High-light effects on CO2
fixation gradients across leaves. Plant, Cell and Environment 19:1261-1271.
SUN J, AND J.N. NISHIO. 2001. Why abaxial illumination limits photosynthetic carbon
fixation in spinach leaves. Plant and Cell Physiology 42: 1–8.
ZAR J. H. 1999. Biostatistical analysis. Prentice Hall, Upper Saddle River, New Jersey,
USA.
41
Table 2.1
Pigment content and leaf thickness in the four leaf types of Galax urceolata tested in this
study. Values represent means of 3 samples (±SE) for chlorophyll a (Chl a), chlorophyll b
(Chl b), total chlorophyll (Total Chl), ratio of chlorophyll a to chlorophyll b (Chl a/b),
anthocyanin, and leaf thickness. Row headings describe the coloration of the
adaxial/abaxial surfaces. Green surfaces contain no visible anthocyanin, red surfaces
contain visible anthocyanin.
42
Table 2.1
Chl b
Chl a
Total Chl Chl a/b Anthocyanin Leaf thickness
2
2
(µmol/cm2)
(mm)
(µg/cm ) (µg/cm ) (µg/cm2)
Green/Green
35.9 a
13.5 a
49.4 a
2.64 a
0.0622 a
0.223 a
(3.8)
(0.90)
(4.2)
(0.074)
(0.062)
(0.007)
Green/Red
35.4 a
15.0 a
50.4 a
2.36 a
11.2 b
0.273 a
(3.3)
(0.93)
(4.2)
(0.099)
(5.9)
(0.022)
a
a
a
a
b
14.9
51.4
2.48
12.2
0.257 a
Red/Green
36.5
(4.2)
(2.3)
(6.4)
(0.12)
(2.8)
(0.0033)
a
a
a
a
c
Red/Red
34.2
14.5
48.6
2.36
23.7
0.273 a
(0.88)
(0.21)
(0.68)
(0.16)
(3.8)
(0.014)
Note: Means within a column followed by different letters are different at p< 0.05.
43
Figure 2.1
Cross sections of leaves exhibiting the range of spatial distribution of anthocyanin seen in
winter leaves of the evergreen herb, Galax urceolata. Depicted here are leaves with: (A)
Acyanic adaxial and abaxial, (B) Anthocyanic adaxial and acyanic abaxial, (C) Acyanic
adaxial and anthocyanic abaxial, and (D) Anthocyanic adaxial and abaxial surfaces. In all
pictures the adaxial surface is facing upwards.
44
Figure 2.1
45
Figure 2.2
Spectral scans of Galax urceolata leaves exhibiting four combinations of pigmented
surfaces: green adaxial and green abaxial (solid line), green adaxial and red abaxial (dotted
line), red adaxial and green abaxial (dashed line), and red adaxial and red abaxial (dashed
and dotted line). Graphs in left column represent (A) Absorptance, (C) Transmittance, and
(E) Reflectance curves of the leaves when light was shone on the lower surface; graphs in
the right column represent (B) Absorptance, (D) Transmittance, and (F) Reflectance
curves of the leaves when light was shone on the upper surface. Lines are means of three
leaves, and the color (black or gray) indicates the color of the surface that received
experimental illumination; black represents green surfaces, gray represents red surfaces.
46
Figure 2.2
47
Figure 2.3
Recovery of maximum PSII efficiency (Fv/Fm) for red and green Galax urceolata leaves.
In A, light was shone on adaxial surfaces only; in B, only abaxial surfaces were illuminated.
White areas represent high stress treatments and gray areas represent recovery periods.
Closed circles represent leaves with green adaxial and abaxial surfaces; open circles, leaves
with red adaxial and green abaxial surfaces; open triangles, leaves with red adaxial and
abaxial surfaces. Individual values are means ± SE of five replicates. Graph A and some
data from Graph B are from Hughes et al., (2005).
48
Figure 2.3
49
CHAPTER III
OPTICAL EFFECTS OF ABAXIAL ANTHOCYANIN ON ABSORPTION OF RED
WAVELENGTHS BY UNDERSTORY SPECIES: REVISITING THE BACK-SCATTER
HYPOTHESIS.
Nicole M. Hughes, Thomas C. Vogelmann, and William K. Smith
The following manuscript was published in Journal of Experimental Botany, volume 59,
pages 3435-3442, 2008, and is reprinted with permission. Stylistic variations are due to the
requirements of the journal. N.M. Hughes performed the laboratory and field work, data
analysis, and prepared the manuscript. T.C. Vogelmann provided ideas, equipment, and
technical assistance, W.K. Smith provided equipment, logistical support, and editorial
assistance.
50
Abstract
Red/purple coloration of lower (abaxial) leaf surfaces is commonly observed in
deeply-shaded understory plants, especially in the tropics. However, the functional
significance of red abaxial coloration, including its role in photosynthetic adaptation,
remains unclear. The objective of this study was to test the back-scatter hypothesis for
abaxial leaf coloration, which posits that red pigments internally reflect/scatter red light
transmitted through the upper leaf surface, thereby enhancing photon capture by the
mesophyll in light-limited environments. Abaxially red/non-red variegated leaves of
Begonia heracleifolia (L.) were used to compare reflectance spectra and chlorophyll
fluorescence profiles of abaxially anthocyanic (red) and acyanic (non-red) tissues under red
light. Photosynthetic gas exchange in response to red light was also compared for abaxially
red/non-red leaf sections. Results did not support a back-scattering function, as
anthocyanic leaf surfaces were not more reflective of red light than acyanic surfaces.
Anthocyanic tissues also did not exhibit increases in mesophyll absorptance of red light, or
increased photosynthetic gas exchange under red light at any intensity, relative to acyanic
tissues. These results suggest that abaxial anthocyanins do not significantly enhance
absorption of red light in the species tested, and alternative functions are discussed.
51
Introduction
Anthocyanins are vacuolar pigments commonly responsible for red, purple, and
blue coloration of plant tissues. They can be found in plant species across a broad range of
habitats, and are especially common in understory plants of the tropics (for species surveys
see Forsyth and Simmonds, 1954; Lee and Collins, 2001; Dominy et al., 2002). Among
tropical species, leaf coloration may be transient—occurring in upper (adaxial) and/or
lower (abaxial) cell layers during development and senescence or, alternatively, coloration
may be permanent (Lee and Collins, 2001). In permanently pigmented leaves of the tropics,
anthocyanins are most commonly abaxial (Lee et al., 1979; Lee and Collins, 2001).
Despite the widespread distribution of abaxial coloration among tropical taxa, very little is
known regarding the functional significance of this trait. Previous studies on the
ecophysiology of foliar anthocyanin have focused almost exclusively on plants with
pigmentation in the upper (adaxial) surface of leaves. In high-light environments, for
example, it has been demonstrated that adaxial pigmentation may function in lightattenuation, protecting underlying cells from photoinhibition through absorption of high
energy blue-green wavelengths (Gould et al., 2002; for reviews see Chalker-Scott, 1999,
Close and Beadle, 2003, and Gould, 2004). Abaxial anthocyanins have also been
implicated to function in photoprotection in plants where exposed abaxial leaf surfaces are
vulnerable to high-incident light (Drumm-Herrel and Mohr, 1985; Sherwin and Farrant,
1998; Hughes and Smith, 2007). However, several studies have also yielded results that do
not support a photoprotective function (e.g. Burger and Edwards, 1996; Dodd et al., 1998;
Lee et al., 2003; and Kyparissis et al., 2007). In addition to functioning in light attenuation,
anthocyanins in upper cell layers have also been suggested to function in insect avoidance
52
and/or deterrence (Stone, 1966; Hamilton and Brown, 2001; Archetti and Brown, 2004;
Lev-Yadun et al., 2004; Karageorgou and Manetas, 2006), act as a fungicide (Coley and
Aide, 1989), function as a carbon sink (Arnold et al., 2004), or as antioxidants during
periods of high photo-oxidative stress (Neill et al., 2002; Neill and Gould, 2003). However,
none of these hypotheses seem appropriate for explaining the occurrence of anthocyanin in
the abaxial cells of understory species.
Smith (1909) proposed the first functional explanation for abaxial coloration in
understory plants, suggesting that anthocyanin pigments function to elevate leaf
temperatures by increasing sunlight absorption, possibly increasing transpiration, nutrient
uptake, metabolism, and growth rates in understory plants. The idea that anthocyanins
increase leaf temperature has not received experimental support, as field measurements
have shown no differences in leaf temperature between abaxially anthocyanic and acyanic
leaves (Lee et al., 1979; Lee et al., 1987; Gould et al. 1995). Lee et al. (1979) observed
that the abaxial leaf surfaces of some abaxially anthocyanic species appeared to reflect
more red light than green surfaces. This observation led to the idea that anthocyanin
pigments near the lower epidermis may function to reflect adaxially-transmitted red light
back into the mesophyll, possibly maximizing capture of red photons in environments
where light is limiting. This “back-scatter” hypothesis has yet to receive experimental
validation beyond these initial observations, though it is still frequently invoked to explain
abaxial coloration in understory plants. In the current study, we employed a cultivated
variety of Begonia heracleifolia (L.) with abaxial anthocyanic/acyanic variegation (Fig. 1),
along with recently-developed, non-destructive optical techniques for measuring light
absorbance within leaf tissues, to test the back-scatter hypothesis. We also compared the
53
light response of photosynthesis under red light for abaxially-anthocyanic and -acyanic leaf
sections to evaluate whether abaxial anthocyanins confer a significant photosynthetic
advantage in leaf tissues.
54
Materials and methods
Plant material
Begonia heracleifolia plants were grown in shaded greenhouse conditions with
periodic exposure to lower intensity, short-lived sunflecks to longer lasting, more intense
sun-patches (Smith et al., 1989). Photosynthetic photon flux densities (PPFDs) in the
shade averaged ca.50 μmol m-2 s-1, while irradiance during sunflecks and sun-patches
ranged from ca. 150 - 1500 μmol m-2 s-1. In all measurements comparing abaxially red and
non-red tissues, adjacent variegated sections of individual leaves were used.
Leaf sections for light microscopy were cut from fresh leaves using a razor blade,
and mounted on a Zeiss Axioplan upright microscope (Carl Zeiss Inc., Thornwood, NY,
USA). Sections were viewed using differential interference contrast (DIC) microscopy,
and images were captured using a Hamamatsu C5810 three-chip cooled color CCD camera
(Hamamatsu Photonics, Hamamatsu City, Japan). Photographs of leaf sections were
rotated and adjusted for brightness and image sharpness using Adobe Photoshop CS
(Adobe Systems, San Jose, CA, USA).
Spectral scans
Five leaves were removed from individual plants, kept under cool, moist conditions,
and analyzed within 8 hrs using a Li-Cor 1800 spectroradiometer with external integrating
sphere (Li-Cor, Inc., Lincoln, Nebraska, USA). Measurements were taken at 2 nm intervals,
and absorbance was calculated as (1 – reflectance – transmittance). Spectral data were
compared by calculating 95% confidence intervals for five replicates; areas of error-bar
overlap were considered not considered to be significantly different. For ambient sunlight
55
measurements, the spectroradiometer was positioned horizontally on a flat surface on a
clear, sunny day in Winston-Salem, NC, USA. For transmission scans of the red optical
filter used in photosynthesis experiments (transmittance spectrum shown in Figure 2)
(Schott Glass, Grünenplan, Germany), the filter was placed in the water bath beneath a
1000W metal halide bulb, and the spectroradiometer was placed in at the same height
below the bulb as the cuvette used in experimentation.
Pigments
Chlorophyll was extracted from the same leaves for which spectral curves had been
measured. Chlorophylls were extracted by removing three 0.33 cm2 leaf discs from either
abaxially anthocyanic or acyanic leaf sections, and extracting the tissues overnight in 3 mL
N, N’-dimethylformamide. Chlorophyll a and b concentrations were determined
spectrophotometrically using the equations described by Porra (2002). Chlorophyll a/b
ratios were calculated to assess relative emphasis on light capture versus processing for red
and green sections, as higher chlorophyll a/b ratios reflect both an increase in core
(exclusively chl a) relative to light-harvesting (both chl a and b) complexes, and/or higher
ratios of Photosystem I (4 / 1 ratio of chl a/b) to Photosystem II (1.2 / 1 ratio) (Cui et al.,
1991; Demmig-Adams, 1998; Hopkins and Hüner, 2004). Pigment concentrations of
adjacent red and green sections were compared using a one-tailed, paired t-test, with p <
0.05 used to determine significance.
56
Chlorophyll fluorescence profiles
Spatial distribution of light absorption within leaf tissues was determined as
described in Vogelmann and Han (2000). Briefly, cm2 sections were cut from six
different variegated leaves, such that one half of each section contained abaxial
anthocyanin and the other half did not (Fig. 1D). The sections were then mounted
on an inverted microscope and upper (adaxial) surfaces were irradiated using filtered blue,
green, and red light (intensity of the beam was ca. 500 μmol m-2 s-1 for blue, 300 for green,
and 200 for red light). Resulting leaf chlorophyll fluorescence was filtered through a
narrow band-pass filter (680 nm, half band width = 16nm, Corion Corp., Holliston, MA),
and captured using a cryogenically cooled charged-coupled device (CCD) camera (CH270
camera head, CF200A 16/40 camera electronics unit, AT200 controller board, 35-mm
shutter; all from Photometrics, Tuscon, AZ). Intensity of chlorophyll fluorescence was
used to quantify light absorbance by chlorophylls (with highest intensities corresponding
with greatest absorbance). Chlorophyll fluorescence according to leaf depth for each image
was quantified using Image Pro Plus software (version 4.1, Media Cybernetics, Silver
Springs, MD). It is important to note that chlorophyll fluorescence in all cases was emitted
exclusively by cells containing chlorophyll (i.e. the mesophyll), though most of this light
appeared to scatter in the adjacent clear epidermal lens cells (as described in Brodersen and
Vogelmann, 2007). Mean percent chlorophyll fluorescence for abaxially anthocyanic and
acyanic tissues was plotted versus leaf depth (from adaxial surface) and 95% confidence
intervals were calculated. Areas of confidence interval overlap were not significantly
different.
57
Light response of photosynthesis
To further test the idea that abaxial anthocyanins enhance capture of red photons for
use in photosynthesis, photosynthetic light response curves were measured for
abaxially anthocyanic/acyanic leaf sections under red light using a Li-Cor 6400
infra-red gas analyzer with clear-top chamber (Licor, Lincoln, Nebraska, USA).
Light was supplemented by a 1000-W metal halide lamp equipped with a UV filter and
water bath, and irradiance was filtered through a red optical filter (transmittance spectrum
shown in Figure 2) (Schott Glass, Grünenplan, Germany). Leaves were positioned
horizontally, with upper surfaces facing the light source. Because variegated leaf sections
were smaller in area than the leaf chamber, an opaque tape with a 2 cm2 square opening
was used to cover the areas of leaves not being measured on both adaxial and abaxial
surfaces. The tape effectively reduced stomatal gas exchange to negligible amounts (i.e.
<1-2%), as determined by experiments on leaf surfaces entirely covered in tape. Neutral
density shade cloths were used to obtain desired PPFDs, which ranged from 0 – 600 µmol
m-2s-1 (the latter representing roughly the maximum intensity of red light in full sunlight;
Lee and Downum, 1991). Quantum yield of photosynthesis was determined as the slope of
the linear portion of the curve, and compared using a paired t-test for adjacent red and
green sections on the same leaf. Sample size for gas exchange experiments was five leaves.
Light response curves for red and green leaf sections were compared using repeated
measures ANOVA.
58
Results and Discussion
Optical effects of abaxial anthocyanin
In contrast to the findings of Lee et al. (1979), red (anthocyanic) abaxial surfaces
of B. heracleifolia were not more reflective of red wavelengths than green (acyanic)
abaxial surfaces, but rather, acyanic leaf surfaces were significantly more reflective of all
wavelengths than anthocyanic (Figs. 3A, B). This was likely due both to the presence of
anthocyanin and higher chlorophyll concentrations in anthocyanic tissues (both of which
increase leaf absorptance, decreasing reflectance) (Table 1, Fig. 3). Increased chlorophyll
content has been observed in several anthocyanic species relative to acyanic phenotypes
(e.g. Gould et al., 1995; Manetas et al., 2003; Hughes et al., 2005), and has been attributed
to shading effects of anthocyanin (discussed in further detail below). Therefore, although
not ideal for quantifying the optical effects of anthocyanin alone (due to confounding
effects of chlorophyll), these scans are likely representative of the realistic effects of
anthocyanin on leaf optics in vivo. Furthermore, important information relevant to the
back-scatter question can still be garnered from these scans. For example, absorbance
spectra showed high absorbance efficiency (~90%) of red light (650-700 nm) in both tissue
types, regardless of the presence of anthocyanin, with less than 5% being lost via
transmittance through the leaf lamina (Fig. 3). This degree of efficiency in photon
absorption is not unique to Begonia, as shade plants are generally known to increase
allocation of resources towards light harvesting capacity, resulting in relatively high
quantum yield of photosynthesis under low irradiances (Larcher, 2003). If abaxial surfaces
of understory plants have evolved to maximize light capture via red-light back-scattering,
they seem to have done so for a narrow waveband that is already being (relatively)
59
efficiently captured. Instead, it would seem that a white reflective layer (which is fairly
common in the plant kingdom) would be of greater benefit than one that reflects red alone,
as this would enhance capture of wavelengths more readily lost through transmittance, such
as green and yellow (Woolley, 1971; Lin and Ehleringer, 1983; DeLucia et al., 1986).
Comparing these scans to those published for other species, the lack of increased
reflectance of red wavelengths in anthocyanic versus acyanic tissues has also been reported
in other studies (Gould et al., 1995; Hughes and Smith, 2007). Slight (1-2%) increases in
reflectance of red wavelengths have also been reported for anthocyanic tissues relative to
acyanic (Burger and Edwards, 1996; Liakopoulos et al., 2006), though differences on the
order of the magnitude of those reported in Lee et al. (1979) (i.e. 20% higher reflectance of
red wavelengths by red tissues) have not been reported. It is possible that repeated
analyses of the species tested by Lee et al. (1979) using newer equipment may yield
different results (D.W. Lee, personal communication).
Effects of abaxial anthocyanin on light capture and photosynthesis
Chlorophyll fluorescence profiles also did not support a back-scattering function of
the abaxial anthocyanin layer. If back-scattering was occurring, increased fluorescence
should have been observed in anthocyanic tissues relative to acyanic under red light,
especially in the most abaxial mesophyll cells. Instead, chlorophyll fluorescence was
similar at all depths in anthocyanic and acyanic leaf sections under red light, with very little
fluorescence observed in abaxial cells layers in either case (Figs. 1A and 4A). This effect
was even more pronounced under blue light (Fig. 1B), where virtually no fluorescence was
observed to be emitted from abaxial mesophyll cells. This indicates that most of the
60
incident red and blue light was absorbed by chloroplasts in the adaxial mesophyll, and that
very little was transmitted to abaxial cells (consistent with our interpretation of spectral
scans). Because the light levels used here (i.e. 200 μmol m-2s-1 for red light) were
substantially more intense than PPFDs reported for the shade of tropical forest understories
(total PPFDs ranging from ca. 2-45 µmol m 2s-1, Lee, 1987), it is unlikely that ambient red
wavelengths in the field would be sufficient to penetrate mesophyll cells unabsorbed, much
less penetrate them, then strike the anthocyanin layer and be back-scattered. Furthermore,
because understory species photosynthetically saturate at relatively low PPFDs (i.e. 200300 μmol m-2s-1; Björkman and Demmig, 1987; Larcher, 2003), irradiance greater than 200
μmol m-2s-1, such as that experienced during longer-lasting, more intense sunflecks or sunpatches, would likely be sufficient to saturate photosynthesis without the assistance of
back-scattering. Consistent with this interpretation, no significant difference in
photosynthesis between abaxially-red and green leaf sections was observed either under
low intensities of red light (0 – 200 μmol m-2s-1; Fig. 5A), or when higher irradiances were
included (0 – 600 μmol m-2s-1; Fig. 5B) (p = 0.73 and 0.68 respectively), even despite
higher chlorophyll content in red tissues. At higher light intensities, some variation in
photosynthetic gas exchange was observed between leaves (most likely due to differences
in stomatal conductance, not shown), though within-leaf comparisons of adjacent red and
green sections consistently showed nearly identical rates of photosynthesis at all intensities
(e.g. Fig. 5C).
61
Alternative functions of the anthocyanic layer
Although no difference in mesophyll absorptance of red light between abaxially red
and green tissues was observed, significant differences in green-light absorption were
apparent in our study. Spectral scans showed that anthocyanic tissues absorbed
significantly more blue-green light (500-600 nm) than acyanic tissues (Fig. 3E),
corresponding with anthocyanin’s absorbance peak. Perhaps more interestingly,
chlorophyll fluorescence profiles of abaxially-red tissues showed significantly reduced
absorption of green light by mesophyll cells (i.e. reduced chlorophyll fluorescence)
compared to acyanic tissues, with effects being greatest in cells most proximate to the
anthocyanic layer (Figs. 1C and 4C). These results are consistent with an alternative
explanation for abaxial coloration proposed by Gould et al. (1995), which purports that
abaxial anthocyanins in understory plants function in light attenuation (similarly to their
function in adaxial cells) during intermittent exposure to high-intensity sunlight (i.e. sunpatches). Gould et al. based this hypothesis on the observation that abaxially-anthocyanic
leaves had significantly higher maximum quantum yield efficiency of photosystem II
(Fv/Fm) than acyanic leaves, as well has higher chlorophyll content and higher maximum
photosynthesis. However, no details were presented to explain how abaxial pigments
might protect leaves in this way.
From the chlorophyll fluorescence profiles presented here, we suspect that
attenuation of green wavelengths by anthocyanins may be occurring (1) as adaxiallyincident light passes through the anthocyanic spongy mesophyll (reducing absorbance by
subjacent chloroplasts), and/or (2) when remaining green light reaches anthocyanic-abaxial
lens cells, being absorbed before internal scattering may occur (i.e. the opposite of back-
62
scattering). This photoprotective function may be especially adaptive in shade-adapted
understory plants, which photosynthetically saturate (and thus photoinhibit) at relatively
low irradiances, and yet frequently encounter potentially damaging irradiance via highintensity sunflecks and sun-patches. The occurrence of the pigment on the abaxial surface
rather than the adaxial might prevent interference of light absorption by anthocyanin at low
light intensities, providing photoprotection only when intensities are high enough to
penetrate the mesophyll (which would likely correspond with damaging levels of photons).
This photoprotective function is further supported by the chlorophyll data (Table 1). As
shown in previous studies on anthocyanic versus acyanic phenotypes (e.g. Gould et al.,
1995, Manetas et al., 2003; Hughes et al., 2005), abaxially-red tissues measured here had
significantly higher total chlorophyll content, and lower chlorophyll a/b ratios than non-red
tissues (p < 0.01 for both). High chlorophyll content and low a/b ratios generally reflect
increased emphasis on light capture relative to processing—a characteristic associated with
more shade-adapted tissues (Demmig-Adams, 1998). These shade-like qualities make
sense for adaxially-anthocyanic tissues (which are directly shaded by the overlying
anthocyanic layer), and also for abaxially-anthocyanic tissues when abaxial attenuation is
considered. However, it is also possible that this effect may be due to increased shading
due to higher chlorophyll content, though this too might be an indirect result of shading by
anthocyanin (as mentioned above).
Conclusion
In summary, the results reported here clearly do not support a back-scattering
function of abaxial red coloration. Red abaxial surfaces were not more reflective of red
63
light than non-red abaxial surfaces, and the presence of a red abaxial layer did not result in
increased mesophyll absorption of red light, or an increase in photosynthetic gas exchange.
However, these results do appear consistent with a photoprotective function of abaxial
anthocyanin, though a more detailed comparison of photoinhibition of photosynthesis for
abaxially anthocyanic versus acyanic tissues is needed to further substantiate this
explanation.
64
Literature Cited
Archetti M, Brown S. 2004. The coevolution theory of autumn colours. Proceedings of the
Royal Society B: Biological Sciences 271, 1219-1223.
Arnold T, Appel H, Patel V, Stocum E, Kavalier A, Schultz J. 2004. Carbohydrate
translocation determines the phenolic content of Populus foliage: a test of the sink– source
of plant defense. New Phytologist 164, 157–164.
Björkman O, Demmig B. 1987. Photon yield of O2 evolution and chlorophyll
fluorescence characteristics at 77 K among vascular plants of diverse origins. Planta 170,
489-904.
Brodersen CR, Vogelmann TC. 2007. Do epidermal lens cells facilitate the absorptance of
diffuse light? American Journal of Botany 94, 1061-1066.
Burger J, Edwards GE. 1996. Photosynthetic efficiency, and photodamage by UV and
visible radiation, in red versus green leaf coleus varieties. Plant Cell Physiology 37, 395399.
Chalker-Scott L. 1999. Environmental significance of anthocyanins in plant stress
responses. Photochemistry and Photobiology 70,1–9.
Close DC, Beadle CL. 2003. The Ecophysiology of Foliar Anthocyanin. The
Botanical Review 69, 149–161.
Coley PD, Aide TM. 1989. Red coloration of tropical young leaves: a possible
antifungal defense? Journal of Tropical Ecology 5, 293-300.
Cui M, Vogelmann TC, Smith WK. 1991. Chlorophyll and light gradients in sun and shade
leaves of Spinacia oleracea. Plant, Cell & Environment 14, 493-500.
DeLucia, EH, Nelson K, Vogelmann TC, Smith WK. 1996. Contribution of intercellular
reflectance to photosynthesis in shade leaves, Plant, Cell & Environment 19, 159-170.
Demmig-Adams B. 1998. Survey of thermal energy dissipation and pigment
composition in sun and shade leaves. Plant Cell Physiology 39, 474–482.
Dodd I, Critchley C, Woodall G, Stewart G. 1998. Photoinhibition in differently coloured
juvenile leaves of Syzygium species. Journal of Experimental Botany 49, 1437-1445.
Dominy NJ, Lucas PW, Ramsden LW, Riba-Hernandez P, Stoner KE, Turner IM. 2002.
Why are young leaves red? Oikos 98, 163-176.
Drumm-Herrel H, Mohr H. 1985. Photosensitivity of seedlings differing in their potential
to synthesize anthocyanin. Physiologia Plantarum 64, 60–66.
65
Forsyth WGC, Simmonds NW. 1954. Survey of the anthocyanins of some tropical plants.
Proceedings of the Royal Society of London. Series B, Biological Sciences 142, 549-564.
Gould KS, Kuhn DN, Lee DW, Oberbauer SF. 1995. Why leaves are sometimes red.
Nature 378, 241–242.
Gould KS, Vogelmann TC, Han T, 1 Clearwater MJ. 2002. Profiles of
photosynthesis within red and green leaves of Quintinia serrata. Physiologia
Plantarum 116, 127-133.
Gould KS. 2004. Nature’s swiss army knife: the diverse protective roles of
anthocyanins in leaves. Journal of Biomedicine and Biotechnology 5, 314-320.
Hamilton WD, Brown SP. 2001. Autumn tree colours as a handicap signal. Proceedings of
the Royal Society: Biological Sciences 268, 1489-1493.
Hopkins WG, Hüner NPA. 2004. Photosynthetic electron transport. In Introduction to
Plant Physiology (eds Hopkins WG,. Hüner NP), pp. 68-71. John Wiley, New York.
Hughes NM, Neufeld HS, Burkey KO. 2005. Functional role of anthocyanins in
high-light winter leaves of the evergreen herb Galax urceolata. New Phytologist 168, 575587.
Hughes NM, Smith WK. 2007. Attenuation of incident light in Galax urceolata
(Diapensiaceae): concerted influence of adaxial and abaxial anthocyanic layers on
photoprotection. American Journal of Botany 94, 784-790.
Karageorgou P, Manetas Y. 2006. The importance of being red when young:
anthocyanins and the protection of Quercus coccifera from insect herbivory and
excess light. Tree Physiology 26, 613-621.
Kyparissis I, Grammatikopoulos G, Manetas Y. 2007. Leaf morphological and
physiological adjustments to the spectrally selective shade imposed by anthocyanins in
Prunus cerasifera. Tree Physiology 27, 849-857.
Larcher W. 2003. The light response of photosynthesis. In Physiological Plant
Ecology (ed W. Larcher), p. 113-114. Springer Publishing, Berlin.
Lee DW, Lowry JB, Stone BC. 1979. Abaxial anthocyanin layer in leaves of
tropical rainforest plants: enhancer of light capture in deep shade. Biotropica 11, 70- 77.
Lee DW. 1987. The spectral distribution of radiation in two neotropical rainforests.
Biotropica 19, 161-166.
Lee DW, Brammeier S, Smith AP. 1987. The selective advantages of anthocyanins in
developing leaves of mango and cacao. Biotropica 19, 40-49.
66
Lee DW, Downum KR. 1991. The spectral distribution of biologically active solar
radiation at Miami, Florida, USA. International Journal of Biometeorology 35, 48-54.
Lee DW, Collins TM. 2001. Phylogenetic and ontogenetic influences on the
distribution of anthocyanins and betacyanins in leaves of tropical plants.
International Journal of Plant Sciences 162, 1141-1153.
Lee DW, O’Keefe J, Holbrook NM, Feild TS. 2003. Pigment dynamics and autumn leaf
senescence in a New England deciduous forest, eastern USA. Ecological Research 18, 677694.
Lev-Yadun S, Dafni A, Flaishman MA, Inbar M, Izhaki I, Katzir G, Ne’eman G. 2004.
Plant coloration undermines herbivorous insect camouflage. BioEssays 26, 1126-1130.
Liakopoulos G, Nikolopoulos D, Klouvatou A, Vekkos K, Manetas Y, Karabourniotis G.
2006. The photoprotective role of epidermal anthocyanins and surface pubescence in young
leaves of grapevine (Vitis vinifera). Annals of Botany 98, 257–265.
Lin ZF, Ehleringer J. 1983. Epidermis effects on the spectral properties of leaves of four
herbaceous species. Physiologia Plantarum 59, 91-94.
Manetas Y, Petropoulou Y, Psaras GK, Drinia A. 2003. Exposed red (anthocyanic) leaves
of Quercus coccifera display shade characteristics. Functional Plant Biology 30, 265-270.
Neill SO, Gould KS, Kilmartin PA, 1 Mitchell KA, Markham KR. 2002.
Antioxidant activities of red versus green leaves in Elatostema rugosum. Plant, Cell &
Environment 25, 539–547.
Neill SO, Gould KS. 2003. Anthocyanins in leaves: light attenuators or antioxidants?
Functional Plant Biology 30, 865-873.
Porra RJ. 2002. The chequered history of the development and use of simultaneous
equations for the accurate determination of chlorophylls a and b. Photosynthesis Research
73, 149-156.
Sherwin H, Farrant JM. 1998. Protection mechanisms against excess light in the
resurrection plants Craterostigma wilmsii and Xerophyta viscosa. Plant Growth Regulation
24, 203–210.
Smith AM. 1909. On the internal temperature of leaves in tropical insolation, with special
reference to the effects of their colour on temperature; also observations on the periodicity
of the appearance of young coloured leaves of trees growing in Peradeniya Gardens.
Annals of the Royal Botanical Gardens, Peradeniya 5, 229-298.
Smith WK, Knapp AK, Reiners WA. 1989. Penumbral effects on sunlight
penetration in plant communities. Ecology 70, 1603-1609.
67
Stone BC. 1966. Pandanus Stickm. in the Malayan Peninsula, Singapore, and lower
Thailand. Malaysian Nature Journal 19, 291-301.
Vogelmann TC, Han T. 2000. Measurement of gradients of absorbed light in
spinach leaves from chlorophyll fluorescence profiles. Plant, Cell & Environment 23,
1303-1311.
Woolley JT.1971. Reflectance and transmittance of light by leaves. Plant Physiology 47,
656-662.
68
Figure 3.1
Top: adaxial view of Begonia heracleifolia leaf. Bottom: chlorophyll fluorescence in
partially red and green variegated leaf sections with upper (adaxial) surface illuminated. In
all pictures, adaxial surface is facing upwards and anthocyanins are present only in the left
half of each photograph, as shown in (D). Images represent chlorophyll fluorescence under
red (A), blue (B), and green (C) light. Note that, while fluorescence is only emitted by
chlorophyll-containing cells, it may also be apparent via scatter within adjacent clear,
epidermal lens cells (see text). Scale bar represents 0.5 mm.
69
Figure 3.1
70
Figure 3.2
Spectral scans. Solid line represents percent mean leaf reflectance for the red abaxial
surface of Begonia heracleifolia leaves. Dotted line represents the transmission spectra for
the red filter used in photosynthesis measurements. The ambient solar spectrum is
represented by the dashed line.
71
Figure 3.2
72
Figure 3.3.
Optical properties (absorptance, transmittance, and reflectance) of the
adaxial and abaxial surfaces of anthocyanic (black lines) and acyanic (grey lines)
tissues of Begonia heracleifolia. Lines represent means of five leaves;
error bars are 95% confidence intervals. The waveband depicted represents the visible
spectrum, with violet-blue wavelengths depicted between 400-500 nm, green 500-580,
yellow-orange 580-630, and red 630-700 nm.
73
Figure 3.3
74
Figure 3.4
Percent chlorophyll fluorescence versus leaf depth for abaxially
anthocyanic (black lines) and acyanic (grey lines) tissues of Begonia heracleifolia
under red, blue, and green light. Error bars represent 95% confidence intervals for six
replicates.
75
Figure 3.4
76
Figure 3.5
Photosynthetic light response curves of abaxially acyanic/green (open circles, dotted lines)
and anthocyanic/red (closed circles, solid lines) leaf tissues under red light. (A) Linear
portion of the light response curves shown in (B); (B) light response curves from 5 leaves
(with each having a single red and green section tested), and (C), a representative pair of
light response curves derived from the same leaf. Error bars represent SE; n = 5.
77
Figure 3.5
78
Table 3.1
Chlorophyll data for abaxially anthocyanic (red) and acyanic (green) portions of Begonia
heracleifolia leaves. Values are means for 5 replicates (± SD). Means within columns not
followed by the same letter are different at p < 0.05.
79
Table 3.1
Abaxial
coloration
Red
Chl a
(μg/cm2)
17.9 a
(0.33)
Chl b
(μg/cm2)
7.97 a
(0.30)
Total Chl
(μg/cm2)
25.8 a
(0.39)
Chl a/b
Green
13.7 b
(1.0)
5.44 b
(0.21)
19.1 b
(1.5)
2.51 b
(0.096)
80
2.25 a
(0.040)
CHAPTER IV
THE COORDINATION OF ANTHOCYANIN DECLINE AND PHOTOSYNTHETIC
MATURATION IN DEVELOPING LEAVES OF THREE DECIDUOUS TREE SPECIES
Nicole M. Hughes, Christianna B. Morley, and William K. Smith
The following manuscript was published in New Phytologist, volume 175, 675-685, and is
reprinted with permission. Stylistic variations are due to the requirements of the journal.
N.M. Hughes performed the laboratory and field work, data analysis, and prepared the
manuscript, C.B. Morley was involved in early organization of the project and ideas, and
W.K. Smith provided equipment, logistical support, and editorial assistance.
81
Abstract
Juvenile leaves in high-light environments commonly appear red due to
anthocyanin pigments, which play a photoprotective role during light-sensitive ontogenetic
stages. The loss of anthocyanin during leaf development presumably corresponds to a
decreased need for photoprotection, as photosynthetic maturation allows leaves to utilize
higher light levels. However, the relationship between photosynthetic development and
anthocyanin decline has yet to be quantitatively described. In the current study,
anthocyanin concentration was measured against photopigment content, lamina thickness,
anatomical development, and photosynthetic CO2 exchange in developing leaves of three
deciduous-tree species. In all species, anthocyanin disappearance corresponded with
development of ~50% mature photopigment concentrations, ~80% lamina thickness, and
differentiation of the mesophyll into palisade and spongy layers. Photosynthetic gas
exchange correlated positively with leaf thickness and chlorophyll content, and negatively
with anthocyanin concentration. Species with more rapid photosynthetic maturation lost
anthocyanin earliest in development. Chlorophyll a/b ratios increased with leaf age, and
were lower than those of acyanic species, consistent with a shading effect of anthocyanin.
These results suggest that anthocyanin re-assimilation is linked closely with chloroplast and
whole-leaf developmental processes, supporting the idea that anthocyanins protect tissues
until light-processing and carbon-fixation have matured to balance energy capture with
utilization.
82
Introduction
One of the most conspicuous developmental changes observed in juvenile leaves as
they mature is color change, with young leaves on new growth tips of many species first
appearing red, purple, pink, or less commonly blue or white, and becoming greener with
leaf age. The red-to-blue coloration of young leaves is most commonly due to the pigment
anthocyanin, appearing within vacuoles of epidermal and/or mesophyll cells within hours
to days of seedling germination, and then decreasing concomitantly with leaf expansion
and maturation (Choinski et al., 2003; Cai et al., 2005). Anthocyanic coloration of juvenile
leaves is nearly ubiquitous in tree species at equatorial latitudes (for species surveys see
Lee & Collins, 2001 and Dominy et al., 2002), where coloration has been observed to
precede and accompany chlorophyll development in species with delayed greening (Kursar
& Coley, 1992; Cai et al., 2005). Expanding leaves of both woody perennials (for species
survey see Price & Sturgess, 1938) and herbaceous species of the temperate zone (see
studies listed in Chalker-Scott, 1999), also commonly exhibit anthocyanin, but during a
more gradual greening process.
Despite its widespread phylogenetic distribution, the functional significance of
anthocyanin in juvenile leaves remains unresolved. Some hypotheses, such as the elevation
of leaf temperature (Smith, 1909), have been clearly dismissed (Lee et al., 1979; Lee et al.,
1987; Hughes, 2004), while others, including camouflage against herbivory (Stone, 1979),
fungicide (Coley & Aide, 1989) or antioxidant activity (Rice-Evans et al., 1997; Wang et
al., 1997), or a signal indicating the presence of unpalatable phenolics (Hamilton & Brown,
2001) have received mixed support (Coley, 1981, 1983; Kursar & Coley, 1992; Dominy et
al., 2002; Neill et al., 2002; Numata et al., 2004; Karageorgou & Manetas, 2006; Manetas,
83
2006). Perhaps the most compelling current explanation for anthocyanin in developing
tissues is that the pigments act as light-attenuators, protecting underlying cells from high
irradiance through absorption of high energy blue-green (and possibly UV) wavelengths of
the solar spectrum (for reviews see Chalker-Scott, 1999 and Gould, 2004). While this latter
function may not be beneficial to species in deeply-shaded habitats, light-attenuation
should be particularly advantageous to species with leaves that initiate maturation at more
sun-exposed, apical tips.
Immature leaves tend to be especially vulnerable to high-light stress due to a
combination of several factors, including immature chloroplast structure (Pettigrew &
Vaughn, 1998; Choinski et al., 2003), reduced capacity and quantity of photosynthetic
enzymes (Miranda et al., 1981a; Pettigrew & Vaughn, 1998), and limited stomatal and
cellular conductance of CO2 (Miranda et al., 1981b; Choinski et al., 2003). As a result,
young leaves growing under high irradiances tend to photosynthetically saturate, as well as
photoinhibit, under substantially lower sunlight levels than mature leaves (Hoflacher &
Bauer, 1982). It is therefore generally beneficial for light capture to be down-regulated
early in leaf development. Observed strategies to decrease light capture in juvenile leaves
include increased xanthophyll pigments (Krause et al., 1995; Barker et al., 1997), inclined
leaf orientation (Jiang et al., 2006), pubescence (Liakopoulos et al., 2006), decreased
chlorophyll content (Choinski et al., 2003; Cai et al., 2005), and light-attenuating layers
such as anthocyanin (Manetas et al., 2002; Karageorgou & Manetas, 2006; Liakopoulos et
al., 2006). Although the physiological benefits of these processes have been addressed and
supported in several studies, little information exists which quantitatively describes the
coordination of photoprotective downregulation with the maturation of the photosynthetic
84
processes which they are assumed to protect. The objective of the present study was to
quantitatively describe the relationship between anthocyanin diminution and photosynthetic
maturation, including development of leaf structure, photopigment accumulation, and
photosynthetic CO2 exchange in three deciduous tree species common in the temperate
zone.
85
Materials and Methods
Pigment composition, lamina developmental characteristics (i.e. thickness and
anatomy), and photosynthetic CO2 exchange were measured for developing leaves of three
deciduous tree species under natural field conditions. These parameters were related to the
disappearance of visible anthocyanin in order to identify possible functional interactions.
Measurements were made at consecutive apical nodes representing increasing stages of
maturation.
Plant material
Species observed in this study were sweetgum (Liquidambar styraciflua L.,
Hamamelidaceae), red maple (Acer rubrum L., Aceraceae), and Eastern redbud (Cercis
canadensis L., Fabaceae) growing along disturbed high sunlight-exposed roadsides in
Winston-Salem, NC, USA (36° 8' N 80° 13' W). For acyanic juvenile leaf comparison of
chlorophyll a/b values, juvenile leaves of English ivy (Hedera helix L., Araliaceae),
forsythia (Forsythia suspensa Thunb., Oleaceae), and Japanese honeysuckle (Lonicera
japonica Thunb., Caprifoliaceae) growing in the same areas were also included. All of
these species continuously produce new growth throughout the summer, and measurements
were taken between July and early September. Since some variation exists in the number
of apical red leaves per branch of a given species, we used branches exhibiting the most
commonly observed number of red leaves per branch in each study: sweetgum branches
most commonly had two apical red leaves; red maple, three; redbud, four (Figure 1).
Leaves were determined to be without anthocyanin when anthocyanin levels were less than
60 μmol m-2, corresponding to values where anthocyanins were no longer visible with the
86
unaided eye. In all experiments, at least five different trees growing in fully-exposed (i.e.
at least 7 h of full sunlight per day) habitats were sampled, using one branch on the east
side of each tree. All trees were saplings < 3 m in height. The most basal leaves of the
same branch on which juvenile leaves were sampled were used to represent “mature”
leaves. Maturity was assumed only if leaf thickness, photosynthesis, and chlorophyll
contents remained constant compared to younger leaves on the branch. All mature leaves
were also exposed to full irradiance (> 1350 µmol m-2 s-1) during much of the day,
including the time of sampling.
Microscopic sectioning
Branches of each species were removed from plants, re-cut underwater, and kept
hydrated until sectioned into 50-100 µm sections using a Vibratome sectioner. Sections
were mounted on a Zeiss Axioplan upright microscope (Carl Zeiss Inc., Thornwood, NY),
viewed using differential interference contrast (DIC) microscopy, and images were
captured using a Hamamatsu C5810 three-chip cooled color CCD camera (Hamamatsu
Photonics; Hamamatsu City, Japan). Photographs of leaf sections were rotated and
adjusted for brightness and image sharpness using Adobe Photoshop CS (Adobe Systems,
San Jose, CA). The backgrounds of all images were also converted to white using Adobe
Photoshop CS.
Leaf thickness
Six whole branches of each species containing the leaf nodes of interest were cut
from trees between 10 : 00 and 11 : 00 h, re-cut underwater, and remained submerged until
87
analysis within 2 h. During this time, no visible wilting was observed to occur, and leaf
thicknesses were not found to significantly differ from non-excised branches (data not
shown). Three to six pieces (depending on variance) of leaf tissue visibly free of
protruding veins were cut out from along the mid-rib of each leaf and immediately
measured using IP54 electronic micrometer calipers with 1µm precision (Fred V. Fowler
Co., Inc, Newton, MA). Average values were compared to those of microscopic sections
measured using a reticle to assure accuracy. No significant difference was observed, as
microscopic values fell within ranges derived using calipers (data not shown). Mean
thickness of each leaf was divided by the average thickness of fully expanded leaves of that
species to estimate percent-mature leaf thickness for each node.
Pigment quantification
Branches were collected between 10 : 00 and 11 : 00 h, during which time all plants
had been exposed to full sunlight for at least 3 h. A standard hole puncher was used to
excise three 0.33 cm2 leaf discs, which were then placed in 3 mL N,N’ -dimethylformamide
to extract in the dark for 24 h. Chlorophyll concentrations were determined
spectrophotometrically using the equations described by Porra (2002), and total carotenoid
concentrations were calculated using equations described by Wellburn (1994). Chlorophyll
concentrations were reported as a function of both volume (using mean leaf thicknesses
calculated for each node, derived from other branches previously measured) and leaf area.
This was done to determine whether increases in chlorophyll content corresponded to
increased leaf thickness, as opposed to increased intracellular chlorophyll (the former being
evidenced by changes in chlorophyll per unit area, but not volume). From the same leaves,
88
three additional leaf discs were excised and placed in 3 mL 6M HCl:H2O:MeOH (7:23:70)
for 24 h at 4°C to extract anthocyanins. Anthocyanin concentration was measured
spectrophotometrically as A530 – 0.24 A653 using an extinction coefficient of 30 000 l mol-1
cm-1 (Murray & Hackett, 1991). All extracts were analyzed using a Hewlett Packard 8453
UV-VIS spectrophotometer (Hewlett Packard, Palo Alto, CA). Concentrations of
chlorophyll and carotenoid pigments for leaves of each node were divided by average
values of fully expanded leaves to estimate the percentage of leaf pigments present relative
to mature leaves.
Chlorophyll a/b ratios were calculated to assess relative emphasis on light capture
versus processing during maturation, as higher chlorophyll a/b ratios reflect both an
increase in core (exclusively chl a) relative to light-harvesting (both chl a and b) complexes,
and/or higher ratios of Photosystem I (4/1 ratio of chl a/b) to Photosystem II (1.2/1 ratio)
(Cui et al., 1991; Demmig-Adams 1998; Hopkins and Hüner, 2004a).
Carotenoid/chlorophyll ratios were also calculated to compare relative photoprotection by
the xanthophylls through leaf development to photosynthetic maturity.
Photosynthetic gas exchange
A portable photosynthesis system (PP Systems Inc. model TPS-1, Amesbury, MA)
was used to measure leaf conductance and photosynthesis for plants in the field.
Measurements were taken between 9 : 00 and 12 : 00 h on plants under clear-sky ambient
sunlight (>1350 µmol m-2 s-1) after the leaf had been enclosed in the cuvette for at least one
minute and oriented directly towards the sun, to allow for light-saturated photosynthesis.
Measured light response curves of all age-class leaves were light-saturated (85% of mean
89
maximum values) at well below (<48%) full sunlight (data not shown), consistent with the
findings of Hoflacher and Bauer (1982) and Wallace and Dunn (1980). Thus, measured
photosynthesis under full sunlight was assumed to represent light-saturated photosynthesis
(Asat) under ambient CO2. The most basal (non-senescent) leaves on the branch were
measured to represent fully mature leaves. Leaf area was measured using a Delta-T leaf
area measurement system (Delta-T Devices Ltd, Cambridge, UK) and photosynthesis was
expressed on a per unit leaf area (projected) and volume basis. Leaf mass was not used to
express photosynthesis due to potential problems associated with variations in mass per
unit volume (e.g. cell wall development).
Statistics
To compare levels of photosynthetic pigments, leaf thickness, and gas exchange of
youngest nodes lacking visible anthocyanin, measurements for the first non-red leaf node
(node 3 for sweetgum, 4 for red maple, and 5 for redbud) were compared using a singlefactor ANOVA with Tukey-Kramer test for means difference. ANOVA with TukeyKramer test was also used to compare these parameters for fully mature leaves. A Pearson
Product Moment Correlation test was used to examine the relationship between leaf node
and pigment concentrations, pigment ratios, leaf thicknesses, and photosynthesis for
individual leaves (as opposed to means). Sample size (N) was five or greater for all
comparisons, with N-1 degrees of freedom. Significance for all tests was determined at p <
0.05.
90
Results
Leaf anatomy
Anthocyanins were either contained within the epidermis (redbud), or mesophyll
cells (red maple and sweetgum) (Figure 1). Red maple and sweetgum contained
significantly more anthocyanin per unit area than redbud in the youngest two leaf nodes (p
< 0.0001) (Figure 2), likely reflecting differences in mesophyll versus epidermal
pigmentation. In red maple and sweetgum, all mesophyll cells initially exhibited reddening,
although anthocyanins eventually became restricted to palisade cells in both species prior
to their disappearance. While reddening was observed to occur in upper (adaxial)
mesophyll cells of all species, the youngest leaves of redbud exhibited intense reddening in
abaxial epidermal cells, which dissipated as leaves matured. Anthocyanins were also
observed in abaxial epidermal cells surrounding protruding leaf veins in redbud and red
maple, even when other abaxial epidermal and/or mesophyll cells contained little or no
anthocyanin.
In all species, anthocyanin did not disappear until after palisade layer differentiation
had begun or was complete (Figure 1). In sweetgum, double palisade layers occurred in the
youngest, expanded leaves observed (apical node 1), and anthocyanin appeared to be
present both during and after palisade formation. In red maple and redbud, anthocyanins
dissipated concomitantly with differentiation and elongation of the palisade layer at nodes
3 and 4, respectively.
Sweetgum leaves were significantly thicker than the other species during early
development (p = 0.022 for node 1, p = 0.0036 node 2, p < 0.0001 node 3), but only
significantly thicker than redbud in later development through maturity, (p < 0.0001 node 4,
91
p < 0.0001 for mature leaves) (Table 1). Thickness of red maple leaves did not
significantly differ from redbud leaves until node 4, after which they became increasingly
thicker (p < 0.0001 for node 4, p < 0.0001 node 5, p = 0.0002 for mature leaves). At the
time of anthocyanin disappearance, lamina thicknesses relative to mature leaves upon
anthocyanin loss did not differ significantly (p = 0.596) between species, and complete
disappearance had occurred by the time leaves were ~80% mature leaf thickness (Figure 2).
Pigment accumulation
In all three species, leaf chlorophyll concentration significantly increased with node
position when expressed on a leaf area basis (r2 = 0.77, 0.68, 0.48 for sweetgum, red maple,
and redbud, respectively; p < 0.0001 in all cases; Table 1 and Figure 2). In contrast,
chlorophyll concentrations did not significantly correlate with leaf node when expressed on
a leaf volume basis (r2 = 0.00, 0.15, 0.14 and p = 0.91, 0.060, 0.061 for sweetgum, red
maple, and redbud, respectively) (Table 1 and Figure 2). All three species had
microscopically visible anthocyanins until mean values of 44% of total chlorophyll per unit
area (relative to mature leaves) had developed. This percentage did not vary significantly
between species (p = 0.262). For chlorophyll expressed on a leaf volume basis, this
percentage was 58% and also did not differ among species (p = 0.182). Anthocyanins were
retained until roughly 49% of mature-leaf carotenoids (per unit area) had developed in all
three species (p = 0.058).
When absolute pigment concentrations were compared (as illustrated in Table 1), as
opposed to percents of maturation, anthocyanin in sweetgum leaves disappeared
significantly earlier (p < 0.01) with regards to chlorophyll content on a unit volume basis
92
compared to red maple and redbud (which did not differ). This translated into 0.797 kg m-3
for sweetgum, and 1.22 and 1.27 kg m-3 for red maple and redbud. On a unit area basis,
anthocyanin loss also occurred significantly earlier in sweetgum compared to red maple (p
= 0.048), but did not significantly differ from redud. On average, sweetgum leaves lost
anthocyanin by the time 138 mg m-2 chlorophyll had developed, while red maple and red
bud retained anthocyanin until 298 and 159 mg m-2 chlorophyll had developed
(respectively). Carotenoids concentration did not significantly vary between species in this
respect (p = 0.19), as anthocyanins had disappeared by the time mean carotenoid levels
reached 63 mg m-2 in all species.
While leaf chlorophyll a/b ratios decreased from the tip-down in acyanic species
(honeysuckle r2 = 0.416, ivy r2 = 0.330, forsythia r2 = 0.561; p < 0.005 for all species), chl
a/b increased in all three anthocyanic species (r2 = 0.76, 0.56, 0.31 for sweetgum, red
maple, and redbud respectively; p < 0.01 for all species) (Figure 2, Table 1). However, the
range of chlorophyll a/b values varied substantially among the three species. Sweetgum
leaves had the broadest range and lowest chlorophyll a/b ratios of all species, ranging
between 1.5 and 3.0 for the nodes examined. Chlorophyll a/b of red maple ranged from 2.4
to 3.3, and from 3.4 to 3.7 in redbud. Carotenoid/chlorophyll ratios significantly decreased
from the tip-down in all anthocyanic species (r2 = 0.79, 0.62, 0.63 for sweetgum, red maple,
and redbud, respectively; p < 0.001 for all species) (Figure 2, Table 1), but of the acyanic
species, only ivy showed significant declines (r2 = 0.56; p < 0.0001; p = 0.18 and 0.12 for
honeysuckle and forsythia respectively; data not shown). Carotenoid/chlorophyll ratios of
the youngest non-red leaf were similar among the anthocyanic species (~108% values of
mature leaves), though a Tukey-Kramer test indicated car/chl ratios of sweetgum to be
93
significantly higher than those of red maple (p = 0.0031), while other species combinations
did not significantly differ. In all anthocyanic species, ranges of carotenoid chlorophyll
ratios of leaf nodes measured were similar (mean of 140% for the youngest leaf node,
100% for the 2nd non-red leaf node).
Gas exchange
The youngest non-red juvenile leaves of sweetgum (third node) had significantly
higher photosynthesis under saturating light conditions (Asat) for both absolute and percentmature values compared to the corresponding values for juvenile leaves of red maple and
redbud (p = 0.0003 for absolute values and 0.0029 for percent mature values per unit area;
p = 0.0018 and 0.002 per unit volume, respectively) (Table 1, Figure 3). Mean Asat (per
unit area) observed for the youngest non-red leaves of sweetgum was 11.5 µmol m-2 s-1
(corresponding to 99% full mature maximum photosynthetic values), while red bud and red
maple averaged 6.5 µmol m-2 s-1 (corresponding to ~60% of mature values). On a per unit
volume basis, Asat for the first non-red leaf of sweetgum was 66.4 mmol m-3 s-1 (130%
mature photosynthesis), 55.5 mmol m-3 s-1 for redbud (77.6% of mature), and 38.1 mmol m3 -1
s for red maple (81.6% of mature). In fully mature leaves, the three species did not
differ significantly with respect to photosynthesis per unit leaf area, which averaged 11
µmol m-2 s-1 (p = 0.275). However, on a per unit volume basis, mature leaves of redbud
had significantly (~33%) higher photosynthesis compared to red maple and sweetgum (p =
0.005). Mean values for mature leaf Asat on a unit volume basis were 72.4 (redbud), 46.7
(red maple), and 50.7 mmol m-3 s-1 (sweetgum). Percent mature photosynthesis in
sweetgum, red maple, and redbud correlated positively to the percent of mature chlorophyll
94
content (area basis) (r2 = 0.53, 0.98, 0.91, and p = 0.001, < 0.0001, < 0.0001 respectively)
and leaf thickness (r2 = 0.86, 0.97, and 0.93; p < 0.0001 for all species), and negatively
correlated with anthocyanin (r2 = 0.90, 0.80, 0.60; p < 0.0001 for all species) (Figure 5).
Though ANOVA tests showed a significant difference between leaf conductance rates of
the three species at the first non-red leaf node (p = 0.0369), Tukey-Kramer test for means
difference did not indicate a significant difference between any two pairs. Mean leaf
conductance values for the youngest non-red leaves of the three species corresponded to
143 (sweetgum), 104 (red maple), and 130 mmol m-2 s-1 (redbud). Similarly, there were no
differences among the three species in the percent of mean maximum leaf conductance
relative to mature leaves at the time of anthocyanin disappearance (p = 0.1850), and the
youngest, fully green leaves of all species averaged 68% of the mean maximum leaf
conductance measured for mature leaves.
95
Discussion
Photopigment accumulation
In all species examined, anthocyanins were visible until approximately 44%
chlorophyll (per unit area), 58% chlorophyll (per unit volume) and 49% of total carotenoid
content (per unit area) had developed (Figure 2). These values did not vary significantly
between species. Similar chlorophyll contents have also been reported for developing
leaves of Rosa sp. and Ricinus communis, in which anthocyanic leaves had up to ~50%
chlorophyll and carotenoid contents of mature leaves (per unit area) (Manetas et al., 2002).
This interspecific similarity in photopigment concentrations observed at the time of
anthocyanin disappearance suggests a level of photosynthetic maturity that no longer
requires photoprotection by anthocyanin. Perhaps photopigment concentrations near 50%
are low enough to avoid excessive light capture (i.e. beyond that which can be efficiently
processed by the immature chloroplast and/or anatomical structure), and/or concentrations
are high enough to contribute significantly to self-shading and (in the case of the
xanthophylls) non-photochemical quenching. Interestingly, these values also correspond
closely with those reported by Lee et al. (2003) during anthocyanin formation in a variety
of temperate deciduous tree species during autumn leaf senescence. In that study,
anthocyanin synthesis was found to be initiated when <50% chlorophyll still remained,
corresponding to <200 mg chl m-2. Here, we report a similar pattern for a different life and
phenological stage, i.e. anthocyanins were present only until ~50% mature chlorophyll
content was reached in maturing leaves, corresponding also to <200 mg m-2.
It is notable that, compared to the more constant increase in chlorophyll per unit
area during leaf development, chlorophyll per unit volume remained relatively unchanged
96
at ~58% of mature leaf values at all measured nodes for all three species (Figure 2; Table
1). This suggests that much of the increase in chlorophyll content per unit area observed
during maturation may be attributable to mesophyll thickening, rather than increased
intracellular chlorophyll content. This is supported by the observations that increases in
chlorophyll per unit area paralleled increases in leaf thickness when plotted against
photosynthetic capacity (Figure 5), and also by the visible elongation of palisade cells
during mesophyll thickening (Figure 1). The maintenance of relatively low chlorophyll
content per unit volume (~58% in young leaves) further supports a decreased emphasis on
light capture in younger leaves, with additional chlorophyll likely being added later in
development following completion of additional leaf-structural and chloroplast maturation
(Smith et al., 1997; Pettigrew & Vaughn, 1998; Niinemets et al., 2004). Although not
significant, it should be noted that chlorophyll per unit volume did increase somewhat
during development in all species (Figure 2), indicating the addition of some intracellular
chlorophyll.
In addition to decreased anthocyanin concentrations during leaf development,
carotenoid/chlorophyll ratios also significantly declined (Figure 2), probably reflecting a
decreased need for photoprotection through non-photochemical quenching (NPQ) as leaves
matured. These results are consistent with findings reported for both acyanic and
anthocyanic juvenile-leafed species (Krause et al., 1995; Barker et al., 1997 respectively),
but contrast results of others reporting no significant change in carotenoid pools during
development, even when juvenile leaves were anthocyanic (Manetas et al., 2002). It has
been suggested that species with elevated carotenoids during juvenile phases may develop
other photoprotective structures when leaves mature (such as highly reflective layers),
97
which effectively replace photoprotection by anthocyanins and/or elevated carotenoid pools
(Manetas et al., 2002). In all species examined here, leaf maturation was accompanied by
gradual whitening (presumably due to the thickening of highly-reflective epicuticular
waxes) on abaxial surfaces (data not shown). These layers likely function to increase
reflectance of incident sunlight on abaxial surfaces of these dorsiventrally asymmetric
leaves, which tend to be more light-sensitive than adaxial surfaces (Sun et al., 1996; Smith
et al. 1997; Sun & Nishio, 2001). Indeed, in redbud and Ailanthus altissima (data not
shown), anthocyanin in the abaxial surface remained visible longer than adaxial
anthocyanin, supporting the importance of abaxial-specific photoprotection in both juvenile
and mature leaves (Hughes & Smith, 2007). The concomitant decline of high
carotenoid/chlorophyll ratios and anthocyanins observed here may therefore represent
another level of coordination leaves exhibit during development— the concerted decline of
juvenile-specific photoprotective strategies with the maturation of others more
characteristic of fully expanded leaves.
Previous studies have demonstrated that developing chloroplasts generally exhibit
higher proportions of chl b relative to chl a in developing leaves, corresponding with higher
proportions of PSII relative to PSI (Pettigrew & Vaughn, 1998). The lag in PSI
development during chloroplast maturation corresponds with decreased unappressed
thylakoid membrane surface area availability early in chloroplast development, which is
where PSI complexes primarily occur (Pettigrew & Vaughn, 1998; Hopkins & Hüner,
2004b). Chl a levels increase as unappressed membranes increase in number. Consistent
with this trend, the anthocyanic species evaluated here had significant increases in
chlorophyll a/b during development (Figures 2 and 4; Table 1). However, these values
98
were much lower than those observed for species with acyanic juvenile leaves, with chl a/b
ratios for two acyanic species actually remaining relatively unchanged during development
(Figure 4). The reduced chlorophyll a/b ratios in red leaves relative to green are likely
attributable to the shading effect of anthocyanin. Previous studies have shown that shade
both retards growth and developmental rates (thus reducing the rate of chl a increase), and
also decreases the ratio of core (exclusively chl a) relative to light-harvesting (both chl a
and b) complexes (Cui et al., 1991; Jones, 1995; Demmig-Adams 1998; Hopkins & Hüner,
2004a). This explanation is supported by the observation that redbud, containing
significantly less anthocyanin in young nodes than the other two species, had the highest
chlorophyll a/b ratios of the three species, presumably due to less shading (Figure 2, Table
1). The shading effect of anthocyanin in juvenile leaves has been investigated in greater
detail by Manetas et al. (2003) using red and non-red variants of Quercus coccifera. In
addition to lower chlorophyll a/b ratios, red juvenile leaves of Q. coccifera had a suite of
other shade characteristics, including thinner leaves and lower levels of xanthophyll cycle
pigments relative to acyanic variants. Presumably, these differences were due to shading
by anthocyanic layers which, as previously mentioned, reduce incoming sunlight via strong
absorption of blue-green light.
Leaf structure and gas exchange
Other factors in addition to photopigment content must also be considered when
evaluating photon absorption and processing, including leaf (mesophyll) thickness,
structural maturation (e.g. spongy and palisade mesophyll differentiation and chloroplast
99
development), leaf conductance of CO2, and photosynthetic CO2 fixation (see Smith et al.,
1997; Evans et al., 2004 for reviews).
A strong relationship was found between leaf thickness and the disappearance of
anthocyanin during leaf maturation. On average, anthocyanin loss occurred only when
~80% of mature leaf thickness had been attained (Figure 2). These values did not
significantly vary between species. Photosynthesis (computed on a leaf area basis) also
showed a strong, positive correlation with leaf thickness (Figure 5), most likely due to
increases in chlorophyll per unit area and chloroplast development (as previously
described), as well as leaf structural maturation.
To infer structural maturation, chlorophyll content expressed per unit volume may
be compared to photosynthesis per unit volume. In red maple and redbud, photosynthesis
expressed per unit leaf volume continued to increase through node five (Figure 3), even
though chlorophyll content per unit volume did not change significantly (Figure 2). This
suggests that either (or both) chloroplast development was still ongoing (in terms of
thylakoid number per granum, membrane complexity, stroma lamellae development, etc.),
or that leaf anatomical structure facilitating light distribution and/or CO2 diffusion was still
in the process of maturation. Although we did not observe chloroplast development in this
study, we were able to observe differentiation of mesophyll into palisade and spongy layers
(Figure 1). These structures can be important for enhancing light propagation and
processing by distributing absorbed light more uniformly throughout the mesophyll (in the
case of palisade cells) and facilitating CO2 diffusion (spongy) (Vogelmann et al., 1996;
Smith et al., 1997). Moreover, these structures were not fully differentiated until the third
leaf node in red maple and redbud (Figure 1). In contrast, sweetgum had more constant
100
(~100% mature values) photosynthesis (expressed per unit volume) between nodes two and
five (Figure 3), with fully differentiated, double palisade layers appearing in the youngest
leaf node with expanded leaves (Figure 1).
Paralleling increases in photosynthesis, leaf conductance of CO2 also increased
significantly during development for all species (Figure 3), with the most rapid increase
between leaf nodes occurring in sweetgum. Because mesophyll conductance is a
significant factor affecting photosynthetic potential, it is to be expected that stomata would
not open fully until leaves are structurally and biochemically mature enough to process
increased internal CO2. This may explain why conductance values in sweetgum
approached mature values earlier than red maple and redbud, corresponding to its more
rapid structural maturity (Figure 1).
The increase in photosynthetic carbon fixation facilitated by a mature internal leaf
anatomy may also provide a greater sink for dissipating excessive sunlight energy,
potentially decreasing the requirement of photoprotective mechanisms such as anthocyanin.
In support of this idea, sweetgum (having the highest photosynthetic rates in young leaves)
lost anthocyanin more quickly during development (i.e. at significantly lower chlorophyll
content) than the other two species examined. By the third leaf node, where leaf
anthocyanin was no longer visible in sweetgum, photosynthesis per unit area had reached
~99% full capacity, although only ~138 mg m-2 (44% of mature) chlorophyll had
developed (Figure 3, Table 1). In contrast, anthocyanin remained visible until the fourth
leaf node in red maple (which had only reached ~60% of full photosynthetic capacity, with
~298 mg m-2 chlorophyll), and the fifth node in redbud (~40% full capacity, and ~159 mg
m-2). This trend of prolonged anthocyanin presence in species with slower photosynthetic
101
maturation is supported further by values reported for C. gummifera (Choinski et al., 2003).
C. gummifera had substantially lower photosynthesis during development relative to the
species here, but also retained anthocyanin until much later in development (i.e. until
~100% full leaf thickness had been obtained and ~400 mg chl m-2 had formed; note,
however, that C. gummifera leaves were substantially thicker than leaves measured here).
The association of rapid structural and photosynthetic maturation with rapid anthocyanin
decline provides further evidence for a strong functional coupling between leaf structure,
anthocyanin presence, and photosynthetic maturation.
102
Literature Cited
Barker DH, Seaton GGR, Robinson SA. 1997. Internal and external photoprotection in
developing leaves of the CAM plant Cotyledon orbiculata. Plant, Cell & Environment 20:
617-624.
Cai Z-Q, Slot M, Fan Z-X. 2005. Leaf development and photosynthetic properties of three
tropical tree species with delayed greening. Photosynthetica 43: 91-98.
Chalker-Scott L. 1999. Environmental significance of anthocyanins in plant stress
responses. Photochemistry and Photobiology 70:1–9.
Choinski JS Jr, Ralph P, Eamus D. 2003. Changes in photosynthesis during leaf expansion
in Corymbia gummifera. Australian Journal of Botany 51: 111–118.
Coley PD. 1981. Ecological and evolutionary responses of tropical trees to herbivory: a
quantitative analysis of grazing damage, plant defenses and growth rates. Dissertation,
Chicago, Illinois, USA: University of Chicago.
Coley, PD. 1983. Herbivory and defensive characteristics of tree species in a lowland
tropical forest. Ecological Monographs 53: 209-233.
Coley PD, Aide TM. 1989. Red coloration of tropical young leaves: a possible antifungal
defence? Journal of Tropical Ecology 5: 293-300.
Cui M, Vogelmann TC, Smith WK. 1991. Chlorophyll and light gradients in sun and shade
leaves of Spinacia oleracea. Plant, Cell & Environment 14: 493-500.
Demmig-Adams B. 1998. Survey of thermal energy dissipation and pigment composition
in sun and shade leaves. Plant Cell Physiology 39: 474–482.
Dillenburg LR, Sullivan JH, Teramura AH. 1995. Leaf expansion and development of
photosynthetic capacity and pigments in Liquidambar styraciflua (Hamamelidaceae)effects of UV-B radiation. American Journal of Botany 82: 878-885.
Dominy NJ, Lucas PW, Ramsden LW, Riba-Hernandez P, Stoner KE, Turner IM. 2002.
Why are young leaves red? Oikos 98: 163-176.
Evans JR, Vogelmann TC, Williams WE, Gorton H. 2004. Light capture by the leaf.
Photosynthetic Adaptation: Chloroplast to the Landscape. New York, NY, USA: Springer,
15-41.
Gould KS. 2004. Nature’s swiss army knife: the diverse protective roles of anthocyanins in
leaves. Journal of Biomedicine and Biotechnology 5: 314-320.
103
Hamilton WD, Brown SP. 2001. Autumn tree colours as a handicap signal. Proceedings of
the Royal Society of London B, Biological Sciences 268:1489-1493.
Hoflacher H, Bauer H. 1982. Light acclimation in leaves of the juvenile and adult life
phases of ivy (Hedera helix). Physiologia Plantarum 49: 366–372.
Hopkins WG, Hüner NPA. 2004a. Photosynthetic electron transport. In: Hopkins WG,
Hüner NP, eds. Introduction to Plant Physiology. New York, USA: John Wiley, 68-71.
Hopkins WG, Hüner NPA. 2004b. Lateral heterogeneity is the unequal distribution of
thylakoid complexes. In: Hopkins WG, Hüner NP, eds. Introduction to Plant Physiology.
New York, USA: John Wiley, 75-76.
Hughes NM. 2004. Functional role of anthocyanins in high light winter leaves of the
evergreen herb Galax urceolata. MSc thesis, Boone, NC, USA: Appalachian State
University.
Hughes NM, Smith WK. 2007. Attenuation of incident light in Galax urceolata
(Diapensiaceae): concerted influence of adaxial and abaxial anthocyanic layers on
photoprotection. American Journal of Botany (in press).
Jiang CD, Gao HY, Zou QI, Jiang GM, Li LH. 2006. Leaf orientation, photorespiration and
xanthophyll cycle protect young soybean leaves against high irradiance in field.
Environmental and Experimental Botany 55: 87-96.
Jones CS. 1995. Does shade prolong juvenile leaf development? A morphological analysis
of leaf shape changes in Cucurbita argyrosperma Subsp. Sororia (Cucurbitaceae).
American Journal of Botany 82: 346-359.
Karageorgou P, Manetas Y. 2006. The importance of being red when young: anthocyanins
and the protection of young leaves of Quercus coccifera from insect herbivory and excess
light. Tree Physiology 26: 613-621.
Krause GH, Virgo A, Winter K. 1995. High susceptibility to photoinhibition of young
leaves of tropical forest trees. Planta 197: 583–591.
Kursar TA, Coley PD. 1992. Delayed greening in tropical leaves: an antiherbivore defense?
Biotropica 24: 256-262.
Lee DW, Lowry JB, Stone BC. 1979. Abaxial anthocyanin layer in leaves of tropical rain
forest plants: enhancer of light capture in deep shade. Biotropica 11: 70-77.
Lee DW, Brammeier S, Smith AP. 1987. The selective advantages of anthocyanins in
developing leaves of mango and cacao. Biotropica 19: 40-49.
104
Lee DW, Collins TM. 2001. Phylogenetic and ontogenetic influences on the distribution of
anthocyanins in betacyanins in leaves of tropical plants. International Journal of Plant
Sciences 162: 1141-1153.
Lee DW, O’Keefe J, Holbrook NM, Feild TS. 2003. Pigment dynamics and autumn leaf
senescence in a New England deciduous forest, eastern USA. Ecological Research 18: 677694.
Liakopoulos G, Nikolopoulos D, Klouvatou A, Vekkos K-A, Manetas Y, Karabourniotis G.
2006. The Photoprotective Role of Epidermal Anthocyanins and Surface Pubescence in
Young Leaves of Grapevine (Vitis vinifera) Annals of Botany 98: 257–265.
Manetas Y, Drinia A, Petropoulou Y. 2002. High contents of anthocyanins in young leaves
are correlated with low pools of xanthophyll cycle components and low risk of
photoinhibition. Photosynthetica 40: 349-354.
Manetas Y, Petropoulou Y, Psaras GK, Drinia A. 2003. Exposed red (anthocyanic) leaves
of Quercus coccifera display shade characteristics. Functional Plant Biology 30: 265-270.
Manetas Y. 2006. Why some leaves are anthocyanic and why most anthocyanic leaves are
red. Flora 201: 163–177.
Miranda V, Baker NR, Long SP. 1981a. Limitation of photosynthesis in different regions
of the Zea mays leaf. New Phytologist 89: 179-190.
Miranda V, Baker NR, Long SP. 1981b. Anatomical variation along the length of the Zea
mays leaf in relation to photosynthesis. New Phytologist 88: 595-605.
Murray JR, Hackett WP. 1991. Dihydroflavonol reductase activity in relation to differential
anthocyanin accumulation in juvenile and mature phase Hedera helix L. Plant Physiology
97: 343-351.
Neill SO, Gould KS, Kilmartin PA, Mitchell KA, Markham KR. 2002. Antioxidant
capacities of green and cyanic leaves in the sun species Quintinia serrata. Functional Plant
Biology 29: 1437-1443.
Niinemets U, Tenhunen, JD, Beyschlag W. 2004. Spatial and age-dependent modification
of photosynthetic capacity in four Mediterrean oak species. Functional Ecology 31: 11791193.
Numata S, Kachi N, Okuda T, Manokaran N. 2004. Delayed greening, leaf expansion, and
damage to sympatric Shorea species in a lowland rain forest. Journal of Plant Research
117: 19-25.
Pettigrew WT, Vaughn KC. 1998. Physiological, structural, and immunological
characterization of leaf and chloroplast development in cotton. Protoplasma 202: 23-37.
105
Porra RJ. 2002. The chequered history of the development and use of simultaneous
equations for the accurate determination of chlorophylls a and b. Photosynthesis Research
73: 149-156.
Price JR, Sturgess VC. 1938. A survey of anthocyanin. VI. Biochemical Journal 32: 16581660.
Rice-Evans CA, Miller NH, Paganga G. 1997. Antioxidant properties of phenolic
compounds. Trends in Plant Science 2: 152-159.
Smith AM. 1909. On the internal temperatures of leaves in tropical insolation, with special
reference to the effect of their colour on temperature; also observations on the periodicity
of the appearance of young coloured leaves of trees growing in Peradinaya Gardens.
Annals of the Royal Botanical Garden. Peradinaya 5: 229-297.
Smith WK, Vogelmann TC, DeLucia EH, Bell DT, Shepherd KA. 1997. Leaf form and
photosynthesis. Bioscience 47: 785-793.
Stone BC. 1979. Protective coloration of young leaves in certain Malaysian palms.
Biotropica 11: 26.
Sun J, Nishio JN, Vogelmann TC. 1996. High-light effects on CO2 fixation gradients across
leaves. Plant, Cell and Environment 19: 1261-1271.
Sun J, Nishio JN. 2001. Why abaxial illumination limits photosynthetic carbon fixation in
spinach leaves. Plant and Cell Physiology 42: 1–8.
Vogelmann, TC, Nishio, JN, Smith WK. 1996. Leaves and light capture: light propagation
and gradients of carbon fixation within leaves. Trends in Plant Science 1: 65-70.
Wallace LL, Dunn EL. 1980. Comparative photosynthesis of three gap phase successional
tree species. Oecologia 45: 331-340.
Wang H, Cao G, Prior RL. 1997. Oxygen radical absorbing capacity of anthocyanins.
Journal of Agricultural Food Chemistry 45: 304-309.
Wellburn AR. 1994. Determination of chlorophyll-a and chlorophyll-b, as well as total
carotenoids, using various solvents with spectrophotometers of different resolution. Journal
of Plant Physiology 144: 307-313.
106
Figure 4.1
Whole branches and microscopic sections of leaves from nodes 1-5 of Liquidambar
styraciflua (sweetgum), Acer rubrum (red maple), and Cercis canadensis (redbud). All
scale bars represent 100µm. In sweetgum, leaf sections were magnified at 100X; red maple,
200X for node 1, 100X for nodes 4-5; redbud, 200X.
107
Figure 4.1
108
Figure 4.2
Pigment concentrations and leaf thicknesses expressed as a percent of mature values.
Shaded areas represent visible presence of anthocyanin. Illustrated are: anthocyanin
content, % chlorophyll content per unit area (closed circles) and per unit volume (open
circles), % chlorophyll a/b, % total carotenoids (per unit area), and % leaf thickness for
Liquidambar styraciflua (sweetgum); Acer rubrum (red maple); and Cercis canadensis
(redbud). Points represent means of five replicates ± SD for pigments and ± SE for lamina
thickness.
109
Figure 4.2
110
Figure 4.3
Percent maximum photosynthesis (Asat) and leaf conductance of water vapor (g) values by
leaf node, expressed as a percent of mature-leaf values. Shaded areas represent the visible
presence of anthocyanin. Illustrated are: percent photosynthesis expressed as per unit area
(closed circles; µmol m-2 s-1) and per unit volume (open circles; mmol m-3s-1) and %
conductance for Liquidambar styraciflua (sweetgum); Acer rubrum (red maple); and
Cercis canadensis (redbud). Points represent means of 5-10 replicates ± SD.
111
Figure 4.3
112
Figure 4.4
Tip-down (nodes 1-5) chlorophyll a/b ratios for species with green developing leaves
(closed symbols) and red (open), compared to mature leaf values. Green (acyanic) species
include Hedera helix (circle), Forsythia suspensa (triangle), and Lonicera japonica
(square). Red species include Acer rubrum (circle), Liquidambar styraciflua (triangle), and
Cercis canadensis (square). Points represent means of 5 replicates ± SD.
113
Figure 4.4
114
Figure 4.5
The relationships between photosynthesis maturation and corresponding changes in
chlorophyll per unit area (thick black), leaf thickness (thick gray), and anthocyanin (thin
black). Sweetgum is represented by the solid lines, red maple by long dashes, and redbud
by dotted lines. Percent of mature photosynthesis in sweetgum, red maple, and redbud
correlated positively to percent of mature chlorophyll content (r2 = 0.53, 0.98, 0.91
respectively) and leaf thickness (r2 = 0.86, 0.97, and 0.93), and correlated negatively to
anthocyanin (r2 = 0.90, 0.80, 0.60). Slopes of these lines were also statistically similar
between species. For chlorophyll content, slopes were 0.179, 0.375, 0.340 for sweetgum,
red maple, and redbud, respectively; for thickness, 0.304, 0.500, 0.393, and for anthocyanin,
-0.750, -0.985, and -0.775.
115
Figure 4.5
116
CHAPTER V
SEASONAL PHOTOSYNTHESIS AND ANTHOCYANIN PRODUCTION IN TEN
BROADLEAF EVERGREEN SPECIES
Nicole M. Hughes and William K. Smith
The following manuscript was published in Functional Plant Biology, volume 34, 10721079, and is reprinted with permission. Stylistic variations are due to the requirements of
the journal. N.M. Hughes performed the laboratory and field work, data analysis, and
prepared the manuscript, W.K. Smith provided equipment, logistical support, and editorial
assistance.
117
Abstract
Leaves of many evergreen species turn red when exposed to high sunlight during
winter due to production of photoprotective anthocyanin pigments, while leaves of other
species, lacking anthocyanin, remain green. Why some species synthesize anthocyanin
pigments while others do not is currently unknown. Furthermore, the relative
photosynthetic performance of anthocyanic (red) and acyanic (green) evergreens has yet to
be described. Here we present seasonal ecophysiological data for five red and green
broadleaf evergreen species. We hypothesize that species which synthesize anthocyanins
in winter leaves correspond to those with the most drastic seasonal photosynthetic declines,
as reduced energy sinks increase vulnerability to photoinhibition and need for
photoprotection. Our results did not support this hypothesis, as gas exchange
measurements showed no difference in mean seasonal photosynthetic capacity between red
and green-leaf species. Consistent with anthocyanin’s shading effect, red-leaf species had
significantly higher chlorophyll content, lower chlorophyll a/b ratios, and higher maximum
light capture efficiency of PSII (Fv/Fm) than green-leaf species during the winter, but not
during the summer (when all leaves were green). We conclude that anthocyanin
production during winter is likely not associated with diminished photosynthetic capacity,
and may simply represent an alternative photoprotective strategy utilized by some species
during winter.
118
Introduction
Evergreen plants exhibit a broad range of adaptations enabling extended
photosynthetic carbon gain during winter months, despite a variety of abiotic stresses (for
reviews see Tranquillini 1964; Nilsen 1992; Uemera and Steponkus 1999; Bigras and
Colombo 2001; Öquist and Huner 2003; Adams et al. 2004). Of particular importance are
adaptations which allow photosynthetic tissues to avoid and/or dissipate excess light energy
(Krause 1994). Because low temperatures inhibit the biochemical reactions of the Calvin
cycle, but do not significantly affect light capture, high-sunlight exposure in combination
with low temperatures may lead to excessive energy captured relative to that used in carbon
fixation (Hüner et al. 1998). This imbalance leads to an increase in energy and electron
transfer to molecular oxygen, production of radical oxygen species, and increased
vulnerability to photo-oxidative damage (Mittler 2002). Adaptations which curtail these
processes generally do so by either reducing photosynthetically active radiation (PAR)
incident on the leaf or chloroplasts (i.e. avoidance strategies) or quenching/dissipating
absorbed energy before transfer to the chlorophyll reaction centers (non-photochemical
quenching). Anthocyanin pigments are an example of an avoidance strategy, as they are
thought to intercept incident blue-green light and dissipate the absorbed quantum energy as
heat, thereby protecting underlying photosynthetic cells from photoinhibition (Lee and
Gould 2002). This shade function has been supported by studies showing reduced
photosynthesis in red-leaf morphotypes of several species relative to green (e.g. Bahler et
al. 1991; Burger and Edwards 1996; Gould et al. 2002) as well as reduced photoinhibition
(e.g. Feild et al. 2001; Hughes et al. 2005; Liakopoulos et al. 2006), often resulting in
119
elevated photosynthesis in anthocyanic leaves relative to acyanic leaves under
photoinhibitory conditions (e.g. Gould et al. 1995; Liakopoulos et al. 2006).
Although the capability for synthesizing anthocyanins is nearly ubiquitous among
angiosperms, not all evergreen angiosperms synthesize anthocyanins in leaves during
winter, when photoprotective mechanisms are generally highly engaged (Adams et al.
2004). Interestingly, many of the evergreen species which do not synthesize anthocyanin
in winter leaves do so in other tissues or during different ontogenetic stages, such as in
juvenile leaves, flowers, stems, roots, senescing leaves, and/or in response to pathogen
infection (see Table 1). Their failure to produce anthocyanins in winter leaves therefore
suggests that anthocyanins are not necessary or beneficial for these species during the
winter season. However, this assumption has not been tested, and no explanation currently
exists for why some broadleaf evergreen species synthesize anthocyanins in winter leaves
while others do not.
One possibility is that anthocyanin synthesis only occurs in species with drastic
seasonal decreases in photosynthetic capacity during the winter, corresponding with an
increased need for photoprotection. A decreased capacity for photosynthesis is known to
render plants intrinsically more vulnerable to photoinhibition, due to a reduction in energy
sinks available for energy dissipation (i.e. photochemical quenching; Osmond 1981;
Powles 1984). This idea has been supported by previous research on anthocyanins in
deciduous tree species, where it has been shown that shade-intolerant species
(characterized by high maximum photosynthesis) tend to not produce anthocyanins during
the autumn, while shade-adapted species (lower maximum photosynthesis) do (Koike
1990; Hoch et al. 2001). Indeed, evergreen species are known to vary greatly in seasonal
120
photosynthetic capacity due to either differences in seasonal growth, and/or general
differences in intrinsic limitations to photosynthesis (e.g. vulnerability to cavitation, waterstress tolerance, freezing damage) (Davis et al. 1999; Uemera and Steponkus 1999;
Verhoeven et al. 1999; Adams et al. 2002; Taneda and Tateno 2005). However, it is
unknown whether such differences correspond with red/green leaf coloration in winter.
To determine whether color change corresponds with any observable patterns in seasonal
photosynthesis, we measured photosynthetic gas exchange, photosynthetic light response
curves, and maximum light capture efficiency of photosystem II (Fv/Fm) for ten broadleaf
evergreen species (representing five different plant families) differing in their production of
anthocyanin during winter.
121
Materials and methods
Sites and species
Species selected for study were mature field plants growing along sun-exposed
road-sides in Jonas Ridge, NC, USA (35° 57’ 20” N, 81° 53’ 55”W; altitude: ca. 1200 m)
on south or southeast-facing sites receiving >6 h full sunlight (i.e. >1350 µmol m-2 s-1 on a
horizontal surface) per day during both summer and winter months. Measurements were
taken on clear sunny days, with little or no cloud cover. Detailed descriptions of the
individual species examined are in Table 1. All hourly time designations are expressed as
Solar Time (List 1971).
Photosynthetic gas exchange
Winter photosynthesis measurements were taken between December 4, 2005 and
March 4, 2006, and December 15 – 17, 2007, and summer measurements on August 2-4,
2007. All measurements were made on first year leaves under full ambient sunlight (>1350
µmol m-2 s-1), and plants were sampled via a standard random walk procedure (Codling and
Hill 2005). Measurements were replicated at times of the day that corresponded to periods
with the highest stomatal conductance. During the summer, these higher conductance
periods occurred in early morning (between 800 and 1100 h) and late afternoon (1600 and
1900 h) when air temperatures were coolest. During the winter, measurements were taken
in mid-afternoon (1100 until 1500 h), when air temperatures were the warmest. A portable
photosynthesis system (PP Systems Inc., model TPS-1, Amesbury, MA, USA) was used to
measure leaf conductance, photosynthetic CO2 assimilation, leaf temperatures, and air
temperature for gas exchange measurements. This instrument was chosen because its small
122
size facilitated the substantial climbing involved in sampling. Measured values were
compared on three different occasions and compared to simultaneous measurements using
a LICOR model Li-6400 (Li-Cor, Inc., Lincoln, NE), and were found to be within ±3%.
Photosynthetic response to light was measured in the field on warm winter days (daytime
maximum air temperatures >17°C) following at least three consecutive days with night
time temperatures >2°C. Measurements were made between 1100 and 1500 h using a LiCor 6400 infrared gas analyzer with red and blue LED light sources (Li-Cor, Inc., Lincoln,
NE). This device was used for light-response curves because the LED transmittance curve
avoids the absorbance spectrum of anthocyanin, allowing photosynthesis to be measured
without the pigment’s shading effect (LED spectrum illustrated in Figure 8-3 of Li-6400
Instruction Manual); this allowed light exposure to be standardized for red and green leaves.
The temperature of the leaf cuvette was set to 20°C during the winter to normalize
measurements between samples, while summer measurements were taken at ambient
temperatures (29 – 35° C). Measurements were initiated at ambient photosynthetic photon
flux densities (PPFD), then increased up to 2000 μmol m−2 s−1, and then decreased
incrementally until irradiance was zero. All measurements were taken on plants that had
been exposed to full sunlight for at least 2 hrs to ensure maximal stomatal opening
(checked by measuring leaf conductance values). The order of individual plant
measurements was randomized each day and sample size was at least n = 3, with different
individuals tested for each species each day.
123
Chlorophyll fluorescence
Maximum light capture efficiency of photosystem II (Fv/Fm) was measured in the
field on mature, current-year leaves between December 4, 2005 and March 4, 2006, and
December 15, 2006 and March 15, 2007, and in summer August 2-4, 2007. Measurements
were randomized according to plant and leaf, and taken between 1100 and 1400h. A PAM
Fluorescence System (Hansatech Institute, model FMS-2, Cambridge, UK) emitting a two
second long, 3 mmol m-2 s-1, 594 nm saturating pulse was used to derive Fv/Fm for all
sampled leaves. Prior to each measurement, plants were dark-adapted for 30 min using
model FMS-2 leaf clips.
Pigment analyses
Chlorophyll concentrations were measured and a/b ratios were calculated to assess
the relative emphasis of light capture versus processing in winter leaves. Increased
chlorophyll content is associated with an emphasis on light capture, as well as increased
chlorophyll a/b ratios, which reflect an increase in core (exclusively chl a) relative to lightharvesting (both chl a and b) complexes, and/or higher ratios of Photosystem I (4/1 ratio of
chl a/b) to Photosystem II (1.2/1 ratio) (Cui et al. 1991; Demmig-Adams 1998; Hopkins
and Hüner 2004).
On February 16, 2007, one mature (fully expanded) and healthy-appearing leaf was
removed from five separate individuals in the field between 1100 and 1300 h, after all
plants had been exposed to full sunlight for at least 3 hrs. A standard hole puncher was
used to excise three 0.33 cm2 leaf discs, which were immediately placed in 3 mL N,N’ dimethylformamide to extract in the dark for 24 hrs. Chlorophyll concentrations were
124
determined spectrophotometrically using the equations described by Porra (2002). From
the same leaves, three additional leaf discs were excised and placed in 3 mL 6M
HCl:H2O:MeOH (7:23:70) for 24 hrs at 4°C to extract anthocyanins. Anthocyanin
concentration was measured spectrophotometrically as A530 – 0.24 A653 using an extinction
coefficient of 30 000 l mol-1 cm-1 (Murray and Hackett 1991). All extracts were analyzed
using a Hewlett Packard 8453 UV-VIS spectrophotometer (Hewlett Packard, Palo Alto,
CA).
Statistics
Linear regression analysis was used to evaluate correlations between temperature
and photosynthesis, leaf conductance, and Fv/Fm, for each species. Slopes and Y-intercepts
of best fit lines for these values were grouped according to the presence or absence of
anthocyanin (i.e. red or green winter leaf color, respectively) and compared using a twosample, two-tailed Student’s t-test (Zar 1999). The effects of leaf color on Fv/Fm,
photosynthesis, and leaf conductance values were analyzed using a nested MANOVA (with
species nested within color), on three winter dates differing in air temperature (Cold: low
temp -9°C, high -3°C; Mild: low -3°C, high 10°C; and Warm: low 8°C, high 20°C; dates
are given in Fig. 2 legend); measurements were also made on one summer day for
comparison (low 31°C, high 18°C). Measurements on different days used random leaves
sampled from multiple (i.e. > 5) plants throughout the plot, invalidating use of the repeated
measures technique (Zar 1999).
For light saturation curves, nested MANOVAs were also used to compare dark
respiration, light-saturated photosynthesis at ambient CO2 (Asat), and photosynthesis (A) at
125
individual irradiance levels for species differing in color. A one-tailed t-test was used to
compare linear slopes of photosynthesis below light saturation for red and green species
during summer and winter. To compare seasonal photosynthetic capacity, summer Asat (i.e.
maximum values observed during the summer) were compared to Asat observed on a warm
winter day (low 8°C, high 20°C, representing the maximum values observed during the
winter) for each species using a one-tailed t-test. A one-tailed Student’s t-test was also
used to compare mean seasonal percent declines in Asat for red and green species.
Photopigment concentrations and chlorophyll a/b ratios of red and green species were
compared using a two-way nested MANOVA, with species nested within leaf color. All
tests employed met distributional and sample size requirements described in Zar (1999),
and significance was accepted at p < 0.05.
126
Results
Winter photosynthesis
Photosynthesis significantly (p < 0.0001) increased with air temperature for all
species during the winter (r2 values for green species: K. latifolia 0.583, R. maximum 0.475,
R. catawbiense 0.711, V. minor 0.627, Rhododendron azalea spp. 0.766; r2 values for red
species: G. urceolata 0.428, L. fontanesiana 0.627, L. japonica 0.772, H. shuttleworthii
0.779, R. azalea 0.857) (Figs 1 & 2). The slope of this increase (m = 0.219) did not
statistically differ between red and green species as a group (p = 0.827).
Leaf conductance was more variable during the winter than photosynthesis, but did
increase significantly with temperature for all species except the green-leafed R. maximum
and V. minor during the winter (p = 0.99 and 0.383 R. maximum and V. minor respectively,
p < 0.05 for all remaining species; Fig. 1). The slope of the increase in conductance
according to temperature during the winter (m = 1.87) did not significantly differ for red
and green species (p = 0.136).
On cold winter sample days, photosynthesis for all plants was at or near zero,
corresponding with near zero leaf conductance values (Fig. 2). There was no significant
difference in Asat or leaf conductance between species differing in leaf color on a cold
winter day (p = 0.303 and 0.754, respectively) or a mild winter day (p = 0.101 and 0.607).
However, on warm days, red-leafed species photosynthesized at significantly higher rates
than green-leafed species (p < 0.0001), and also had significantly higher leaf conductance
(p < 0.0001). However, there was also a significant species effect on warm days (p <
0.0001), with very high Asat and g values for the red-leaf L. japonica, and very low values
127
for the green-leaf V. minor driving statistical differences between groups; values for other
species generally overlapped (Fig. 2).
For light-response curves (Fig. 3), red species had significantly higher Asat than
green species during the winter (p = 0.01), although, again, there were highly significant
interspecific differences (p < 0.0001), and with the exception of L. japonica and red R.
azalea spp., Asat for red and green-leaf species was comparable. Similarly, there was no
difference in Asat between red and green-leaf species during the summer when all leaves
were green (p = 0.75). During winter, red-leaf species also showed significantly higher A
than green species at low PAR values (i.e. < 500 µmol m-2 s-1) (p < 0.01 at 200 and 100
µmol m-2 s-1), and had significantly greater quantum yield of photosynthesis at low PAR
(as evidenced by steeper slopes below the light saturation point) (p = 0.02, where mean m
for red species = 0.016, and green species = 0.0098); the groups did not differ in any of
these parameters during the summer (p > 0.50). During both summer and winter seasons,
dark respiration rates of red and green-leaf species did not differ (p > 0.30).
Seasonal differences in photosynthetic gas exchange
While winter photosynthesis of all species was significantly less than summer
values on both cold and mild days (p < 0.0001 for both), on the warmer days (air
temperature > 16°C) some species were able to photosynthesize at rates that did not
significantly differ from summer rates (green species: V. minor p = 0.09; red species: G.
urceolata p = 0.52; Fig. 4). The remaining species photosynthesized significantly below
summer levels on warm winter days (p < 0.01). As a group, the mean percent seasonal
decline in photosynthesis relative to summer values exhibited by red leaves (30%) did not
128
significantly differ from that of green species (41%) (p = 0.53). At all temperatures
measured, all ten species exhibited significantly reduced conductance during the winter
relative to the summer (p < 0.0001).
Fluorescence
Winter Fv/Fm values increased significantly (p < 0.0001) with temperature for all
species (Fig. 1; r2 values > 0.56 for all green species; r2 values > 0.41 for all red species).
Fv/Fm increased with temperature along the same slope (m = 0.0158) for both red and green
groups (p = 0.733). Fv/Fm values of red species were consistently higher (~0.15 on
average) than those of green species across the temperature range measured, and Yintercepts of regressions were significantly higher for red species as a group compared to
green species (p = 0.016). On cold, mild, and warm winter days, red-leaf species had
consistently higher (~0.15 units on average) mean Fv/Fm than green species (p < 0.0001 for
all; Figs 1 & 2).
Pigments
Species with red winter leaves had significantly higher total chlorophyll (24% on
average), and anthocyanin content than green species during the winter (p < 0.0001 for
both; Table 2). Red species also had significantly (27%) lower chlorophyll a/b ratios than
green species on average (p < 0.0001; Table 2).
129
Discussion
The hypothesis that species with red winter leaves would be characterized by lower
photosynthetic capacity during winter than green-leaf species was not supported in this
study. Mean maximum photosynthesis of red-leaf species during winter did not differ from
green-leaf species on cold, mild, or relatively warm winter days (Figs 1 & 2). In fact, the
two species with the highest photosynthesis values during winter were both red-leaf species
(L. japonica and the red R. azalea sp.). Mean winter declines in photosynthesis relative to
summer values (Fig. 4) also did not significantly differ between red-leaf species relative to
green-leaf species, with some members of both groups having Asat on warm winter days
near values observed during the summer, and others exhibiting more drastic declines (i.e. <
50% summer photosynthesis).
Fluorescence measurements of PSII were consistent with a photoprotective function
by anthocyanin. Red-leaf species had consistently higher maximum light-capture
efficiency of PSII (Fv/Fm) than green-leaf species across all measured temperatures during
winter days (Figs 1 & 2); similar trends were also observed for pre-dawn measurements
during the winter (data not shown). Because Fv/Fm is correlated with sustained nonphotochemical quenching (with lower values corresponding with increased retention of
photoprotective antheraxanthin and zeaxanthin), these results suggest that red-leaf species
rely less on sustained xanthophyll-mediated photoprotection than green-leaf species during
the winter, most likely due to anthocyanin’s attenuation of absorbed sunlight reducing the
need for additional photoprotection (Demmig-Adams et al. 1996). However, it is also
possible that red-leaf species are utilizing more the rapidly-reversible component of NPQ
(i.e. pH-dependent conversion of violaxanthin into antheraxanthin and zeaxanthin), which
130
is lost during dark adaptation; this is unlikely, however, because ΦPSII (non-dark adapted
quantum efficiency of PSII) was also consistently higher in red species than green (data not
shown), by roughly the same order of magnitude as Fv/Fm.
Anthocyanin’s shading effect was also evidenced in the relatively shade-adapted
light response curves of red compared to green leaves (Fig. 3). Under red and blue LED
light (which does not overlap with the absorbance spectrum of anthocyanin), red-leaf
species had significantly steeper slopes for the linear part of the light response curves than
green-leaf species, and higher photosynthesis under low PAR (i.e. < 500 µmol m-2 s-1)
accordingly. The two groups did not differ in these respects during the summer, when all
leaves were green. Higher photosynthesis at low PAR is a common feature of more shadeadapted plants, reflecting either decreased respiration rates and/or a greater emphasis on
light capture relative to light processing (Larcher 2003). Because dark respiration values
were not significantly different between the red-and green-leaf groups during the winter,
increased Asat at low PAR was likely due to greater light capture capacity. Consistent with
this explanation, total chlorophyll content was significantly higher and chlorophyll a:b
ratios significantly lower in red-leaf species compared to green (similar to results reported
in Manetas et al. 2003, Lee et al. 2003, Hughes et al. 2005; Table 2).
Higher photosynthesis at lower light levels may also explain why red-leaf species
were able to photosynthesize at saturated values under ambient sunlight, despite the
shading effect of anthocyanin. Pietrini and Massacci (1998) estimated that anthocyanins at
concentrations near the maximum values reported here (i.e. ~4.4 mol/g) may attenuate
~43% incoming PAR in vivo. Thus, under full sunlight, the amount of unabsorbed
irradiance remaining should still be high enough to saturate photosynthesis (as all red-leaf
131
species photosynthetically saturated near 25% full sunlight; Fig. 3). Perhaps anthocyanins
reduce PAR transmittance sufficiently to curtail photoinhibition, but still transmit enough
to allow the leaf to attain light saturation of photosynthesis, maximizing sunlight capture
and utilization during the winter.
Because red and green-leaf species measured here did not differ in seasonal
photosynthesis, we conclude that red coloration in winter leaves does not appear to
correspond with diminished photosynthetic capacity. Instead, the comparable seasonal
photosynthesis of red leaves compared to green suggests that photoprotection by
anthocyanin represents an alternative (rather than additive) photoprotective strategy
employed by some plants during the winter. Perhaps up-regulation of anthocyanin
compensates for species-specific deficiencies in other means of photoprotection (e.g. NPQ,
antioxidants). Research is currently underway by the authors to investigate this possibility.
Alternatively, it is also possible that red-leaf species are indeed limited in photosynthetic
capacity during the winter, but are able to photosynthesize at rates similar to green species
due to anthocyanin’s photoprotective effect. However, this would be difficult to determine
without, for example, utilizing anthocyanin-less mutants of red-leaf species.
132
Literature Cited
Adams WW III, Demmig-Adams B, Rosenstiel TN, Brightwell AK, Ebbert V (2002)
Photosynthesis and photoprotection in overwintering plants. Plant Biology 4, 545-557.
Adams WW III, Zarter CR, Ebbert V, Demmig-Adams B (2004) Photoprotective strategies
of overwintering evergreens. Bioscience 54, 41-49.
Bahler BD, Steffen KL, Orzolek MD. (1991) Morphological and biochemical comparison
of a purple-leafed and a green-leafed pepper cultivar. Horticultural Science 26, 736.
Bigras FJ, Colombo SJ (2001) Conifer Cold Hardiness. Tree Physiology. Kluwer Academic
Publishers, the Netherlands.
Burger J, Edwards GE. (1996) Photosynthetic efficiency, and photodamage by UV and
visible radiation, in red versus green leaf coleus varieties. Plant Cell Physiology 37, 395399.
Codling EA, Hill NA (2005) Sampling rate effects on measurements of correlated and
biased random walks. Journal of Theoretical Biology 233, 573-588.
Cui M, Vogelmann TC, Smith WK (1991) Chlorophyll and light gradients in sun and shade
leaves of Spinacia oleracea. Plant, Cell & Environment 14, 493-500.
Davis SD, Sperry JS, Hacke UG (1999) The relationship between xylem conduit diameter
and cavitation caused by freezing. American Journal of Botany 86, 1367-1372.
Demmig-Adams B, Adams WWIII, Barker DH, Logan BA, Bowling DR, Verhoeven AS
(1996) Using chlorophyll fluorescence to assess the fraction of absorbed light allocated to
thermal dissipation of excess excitation. Physiologia Plantarum 98, 253-264.
Demmig-Adams B (1998) Survey of thermal energy dissipation and pigment composition
in sun and shade leaves. Plant Cell Physiology 39, 474–482.
Feild TS, Lee DW, Holbrook NM (2001) Why leaves turn red in autumn. The role of
anthocyanins in senescing leaves of red-osier dogwood. Plant Physiology 127: 566-574.
Gould KS, Kuhn DN, Lee DW, Oberbauer SF (1995) Why leaves are sometimes red.
Nature 378, 241-242.
Gould KS, Vogelmann TC, Han T, Clearwater MJ. (2002) Profiles of photosynthesis
within red and green leaves of Quintinia serrata. Physiologia Plantarum 116, 127-133.
Hoch WA, Zeldin EL, McCown BH (2001) Physiological significance of
anthocyanins during autumnal leaf senescence. Tree Physiology 21, 1-8.
133
Hopkins WG, Hüner NPA (2004) Photosynthetic electron transport. Introduction to Plant
Physiology (eds. WG Hopkins, NP Hüner). pp. 68-71. John Wiley, New York.
Hughes NM, Burkey KO, Neufeld HS (2005) Functional role of anthocyanins in high-light
winter leaves of the evergreen herb, Galax urceolata. New Phytologist 168, 575-587.
Hüner NPA, Öquist G, Sarhan F (1998) Energy balance and acclimation to light and cold.
Trends in Plant Science 3, 224–230.
Koike T (1990) Autumn coloring, photosynthetic performance and leaf development of
deciduous broad-leaved trees in relation to forest succession. Tree Physiology 7, 21-32.
Krause GH (1994) Photoinhibition induced by low temperatures. Photoinhibition of
Photosynthesis. From Molecular Mechanisms to the Field (eds. NR Baker, JR Bowyer). pp.
331—348. BIOS Scientific Publ., Oxford.
Larcher W (2003) The light response of photosynthesis. Physiological Plant Ecology. pp.
111-120. Springer, NewYork.
Lee DW, O’Keefe J, Holbrook NM, Feild TS (2003) Pigment dynamics and autumn leaf
senescence in a New England deciduous forest, eastern USA. Ecological Research 18, 677694.
Lee DW, Gould KS (2002) Why leaves turn red. American Scientist 90, 524-531.
Liakopoulos G, Nikolopoulos D, Klouvatou A, Vekkos K-A, Manetas Y, Karabourniotis G.
2006. The photoprotective role of epidermal anthocyanins and surface pubescence in young
leaves of grapevine (Vitis vinifera). Annals of Botany 98, 257–265.
List RJ (1971) Smithsonian Meteorological Tables. 527 pp. Smithsonian Institution Press,
Washington D.C.
Manetas Y, Petropoulou Y, Psaras GK, Drinia A (2003) Exposed red (anthocyanic) leaves
of Quercus coccifera display shade characteristics. Functional Plant Biology 30, 265-270.
Mittler R (2002) Oxidative stress, antioxidants, and stress tolerance. Trends in Plant
Science 7, 405-410.
Murray JR, Hackett WP (1991) Dihydroflavonol reductase activity in relation to
differential anthocyanin accumulation in juvenile and mature phase Hedera helix L. Plant
Physiology 97, 343-351.
Nilsen ET (1992) Thermonastic leaf movements: a synthesis of research with
Rhododendron. Botanical Journal of the Linnean Society 110, 205-233.
Osmond CB (1981) Photorespiration and photoinhibition; some implications for the
energetics of photosynthesis. Biochemica et Biophysica Acta 639, 77-98.
134
Öquist G, Hüner NPA (2003) Photosynthesis of overwintering evergreen plants. Annual
Review of Plant Biology 54, 329–355.
Pietrini F, Massacci A (1998) Leaf anthocyanin content changes in Zea mays L. grown at
low temperature: Significance for the relationship between the quantum of PS II and the
apparent quantum yield of CO2 assimilation. Photosynthesis Research 58, 213-219.
Porra RJ (2002) The chequered history of the development and use of simultaneous
equations for the accurate determination of chlorophylls a and b. Photosynthesis Research
73, 149-156.
Powles SB (1984) Photoinhibition of photosynthesis unduced by visible light. Annual
Review of Plant Physiology 35, 14-44.
Taneda H, Tateno M (2005) Hydraulic conductivity, photosynthesis and leaf water balance
in six evergreen woody species from fall to winter. Tree Physiology 25, 299-306.
Tranquillini W (1964) The physiology of plants at high altitudes. Annual Review of Plant
Physiology 15, 345-362.
Uemera M, Steponkus PL (1999) Cold acclimation in plants: relationship between the lipid
composition and the cryostability of the plasma membrane. Journal of Plant Research 112,
245-254.
Verhoeven AS, Adams WWIII, Demmig-Adams B (1999). The xanthophyll cycle and
acclimation of Pinus ponderosa and Malva neglecta to winter stress. Oecologia 118, 277287.
Zar JH (1999) Biostatistical analysis. Prentice Hall, Upper Saddle River, New Jersey, USA.
135
Table 5.1
Detailed descriptions of species used in this study. An asterisk by species name denotes
species which synthesize anthocyanin in winter leaves
136
Table 5.1
Plant form
Origins
Anthocyanic tissues
Species
Family
*Galax urceolata (Poir.)
Brummitt
Diapensiaceae
Forb/herb
Native
Winter rhizomes and
petioles, winter, juvenile
and injured leaves
*Leucothoe fontanesiana
(Steud.) Sleumer
Ericaceae
Shrub
Native
Winter stems, winter,
juvenile, and injured leaves
*Lonicera japonica (Thunb.)
Caprifoliaceae
Vine
Introduced
*Hexastylis shuttleworthii
(Britten & Baker)
Aristolochiaceae
Forb/herb
Native
Winter leaves, winter
petioles, flowers
*Rhododendron spp.
(horticultural azalea)
Ericaceae
Shrub
Introduced
Flowers, new growth stems
senescing leaves
Ericaceae
Shrub
Native
Juvenile and injured leaves,
stems, flowers,
Ericaceae
Shrub
Native
Apocynaceae
Vine/
groundcover
Introduced
Ericaceae
Shrub
Native
Ericaceae
Shrub
Introduced
Runners, winter leaves
Kalmia latifolia (L.)
Rhododendron maximum (L.)
Glands on juvenile leaves
and stems, developing
stems, flowers
Vinca minor (L.)
Flowers
Rhododendron catawbiense
(Michx.)
Flowers, senescing leaves
Rhododendron spp.
(horticultural azalea)
137
Flowers, new growth stems
senescing leaves
Table 5.2
Winter photosynthetic and non-photosynthetic pigment content, expressed per gram of
fresh weight. Gaultheria procumbens was used as a red representative instead of H.
shuttleworthii due to limited availability of H. shuttleworthii leaves for destructive analyses.
138
Table 5.2
Total Chl
(mg/g)
Chl a/b Anthocyanin
(mol/g)
K. latifolia
0.886
(0.22)
4.63
(0.88)
-
R. azalea spp
0.810
(0.20)
4.45
(0.62)
-
R. catawbiense
0.987
(0.13)
4.29
(0.75)
-
0.793
(0.16)
3.61
(0.34)
-
R. maximum
0.857
(0.14)
2.11
(0.30)
-
Red spp. L. fontanesiana
1.08
(0.29)
3.28
(0.34)
1.04
(0.32)
L. japonica
0.623
(0.087)
3.10
(0.30)
2.61
(0.34)
G. urceolata
1.61
(0.26)
2.21
(0.33)
3.27
(1.0)
R. azalea spp
0.978
(0.19)
2.29
(0.28)
5.3
(1.4)
G. procumben
1.10
(0.11)
3.07
(0.71)
1.05
(0.55)
Green spp.
V. mino
139
Figure 5.1
Winter maximum light capture efficiency of PSII (Fv/Fm), light-saturated photosynthesis
(Asat), and leaf conductance (g) for red (dashed lines) and green (solid lines) broadleaf
evergreen species measured across a range of ambient temperatures.
Left column: representative green species (K. latifolia), middle column: representative red
species (G. urceolata), right column: best fit lines of all red (dashed) and green (solid)
species measured. Red species: L. japonica (A), R. azalea spp. (B), G. urceolata (C), L.
fontanesiana (D), H. shuttleworthii (E). Green species: R. catawbiense (F), K. latifolia (G),
R. azalea spp. (H), R. maximum (I), V. minor (J).
140
Figure 5.1
141
Figure 5.2
Cold, mild, and warm winter day measurements of Asat, g, and Fv/Fm for green-leaf and
red-leaf species. Bars represent means of 5-15 replicates ± SD. Cold winter day
measurements were taken on January 29, 2007 (low -9°C , high temp -3°C); mild, February
7, 2006 (low -3°C, high 10°C); warm, December 15, 2006 (low 8°C, high 20°C); summer
measurements taken on June 8, 2006 (low 12°C, high 23°C).
142
Figure 5.2
143
Figure 5.3
Seasonal light response curves for red and green broadleaf evergreen species. Top row:
summer; bottom row: winter. Left column: representative green species (K. latifolia),
middle column: representative red species (G. urceolata), right column: best fit lines of all
red (dashed) and green (solid) species measured. Red species: L. japonica (A), R. azalea
spp. (B), G. urceolata (C), L. fontanesiana (D), H. shuttleworthii (E). Green species: R.
catawbiense (F), K. latifolia (G), R. azalea spp. (H), R. maximum (I), V. minor (J).
144
Figure 5.3
145
Figure 5.4
Percent decrease in maximum observed winter Asat relative to summer values.
White bars represent green species, grey bars represent red species. Asterisks denote
significant declines in maximum winter photosynthesis relative to summer values (p <
0.05).
146
Figure 5.4
147
CHAPTER VI
CONCLUSIONS AND FUTURE DIRECTIONS
148
The results presented here are generally consistent with a photoprotective
explanation of anthocyanin pigments in photosynthetic tissues; however, unresolved issues
still remain. As described in Chapters II and III, abaxial anthocyanin pigments were
demonstrated to be of functional importance in photoprotection. In the case of high-light
species with vertical leaf orientations and/or in environments characterized by high
substrate albedo (Galax urceolata in Chapter II), the abaxial anthocyanic layer directly
intercepted irradiance incident on the more light-sensitive abaxial leaf surface, effectively
reducing photoinhibition of photosynthesis. In species adapted to the deep shade (Begonia
heracleifolia in Chapter III), abaxial anthocyanins also appeared to function in
photoprotection, though not through direct interception of incident light, but rather, through
attenuation of internally-scattered green light transmitted through the adaxial leaf surface.
This latter adaptation would presumably function to buffer internal light levels during brief
to extended periods of high light (e.g. sunflecks or sun-patches) in species adapted to the
deep shade, without interfering with light interception during periods of low light (as
adaxial anthocyanins might). Despite the experimental support drawn from these studies
on abaxial anthocyanin, however, there still remain cases of abaxial pigmentation for which
neither of these explanations seems appropriate. For example, some species of water lilies
exhibit abaxial coloration in floating leaves. Clearly, because of their horizontal orientation
against the water surface, abaxial surfaces of water lilies are not directly exposed to high
light (as was the case with G. urceolata), and so direct interception of incident light seems
an improbable function. Furthermore, these species are almost continuously exposed to
high adaxial irradiance, and so, adaxial anthocyanins would seem more appropriate for
photoprotection than abaxial (in contrast with understory species). There is currently no
149
adaptive explanation for abaxial coloration in these floating leaves. Another interesting,
yet unresolved, case of abaxial pigmentation is that of abaxial red/green variegation (e.g.
Figure 3.1), which is usually observed in understory plants. Although an abaxially
red/green variegated plant was used in Chapter III to test the photoprotection hypothesis, it
is still unclear why this species (and others) would evolve pigmentation in only portions of
leaf tissues. Herbivory avoidance seems one likely explanation, as leaves may appear
smaller or damaged compared to fully green leaves. This hypothesis, however, has not yet
been tested. It is also worth noting that in Begonia heracleifolia (Figure 3.1) it is the
tissues most distal to the major veins that exhibit pigmentation. Perhaps the increased
vulnerability to dehydration stress exacerbates oxidative damage, rendering distal tissues in
greater need of photoprotection. This hypothesis too has not been tested.
In Chapter IV, the coordination between anthocyanin decline and photosynthetic
maturation (and thus, decreased need for photoprotection) was demonstrated using
developing leaves of three deciduous-tree species. This was the first study to
quantitatively describe the association between relative need for photoprotection and
anthocyanin concentration. It should be noted, however, that this relationship was purely
correlative, and could also be attributed to other functions of anthocyanin in addition to
photoprotection. For example, because young leaves are more vulnerable to herbivory
than mature leaves (due to a thinner lamina and cuticle), red coloration may function as a
camouflage to insects lacking a red photoreceptor. This explanation is consistent with the
correlation between loss of redness and attainment of 80% leaf thickness, though it does
not necessarily explain the correlation with development of leaf structural anatomy or
photopigments. Additionally, juvenile leaves might also require anthocyanin as an
150
antioxidant during development, for the same reasons it would be beneficial as a lightattenuating molecule (i.e. increased vulnerability to photo-oxidative damage during
development). This study did not discern whether anthocyanins were functioning as one
or the other, but rather, simply correlated their presence with need for photoprotection.
Future studies should aim to discern between these two functions.
In the study presented in Chapter V, it was hypothesized that red-leafed species
required anthocyanins because of intrinsic limitations to photosynthetic capacity during
winter, which would increase need for photoprotection accordingly. However, results
showed that red-leafed species did not differ in photosynthetic capacity relative to greenleafed species during either summer or winter. Anthocyanins did appear to function in
photoprotection though, as red-leafed species exhibited significantly reduced sustained
photoinhibition of photosynthesis compared to green-leafed species, and also showed
symptoms of shade adaptation. Regardless, it remains unclear why certain evergreen
species would synthesize anthocyanin in winter leaves while others would not. One
possibility is that red-leaf species are indeed limited in photosynthetic capacity during the
winter, but are able to photosynthesize at rates similar to green species due to
anthocyanin’s photoprotective effect. However, this would be difficult to determine
without, for example, utilizing anthocyanin-less mutants of red-leaf species. Another
possibility is that anthocyanin simply represent an alternative (rather than additive)
photoprotective strategy employed by some plant species during the winter, perhaps
compensating for species-specific deficiencies in other modes of photoprotection (e.g. NPQ,
antioxidants). I have begun to test this hypothesis in a study not presented in this
dissertation, and have thus far found no significant difference in low molecular weight
151
antioxidants between red and green-leafed species; xanthophyll analyses are currently
underway. Finally, a third possibility is that anthocyanins are synthesized to alleviate
oxidative stress resulting from low water potentials, either through acting as an antioxidant
or through light-attenuation. According to this hypothesis, species which synthesize
anthocyanin during winter would correspond with those exhibiting the most extreme
declines in daily water potentials (i.e. drought tolerators). This experiment is also currently
underway.
In conclusion, although unanswered questions remain regarding the photoprotective
role of anthocyanins pigments in photosynthetic tissues, the studies presented in this
dissertation were successful in strengthening the evidence for an important photoprotective
function.
152
Nicole Michelle Hughes
Wake Forest University
Department of Biology
Winston-Salem, NC 27109
(336) 414-4116
Email: [email protected]
EDUCATION
B.S. Biology and Education, 2001, Stetson University
M.S. Biology, 2004, Appalachian State University
Advisor: Dr. Howard S. Neufeld
Ph.D. Candidate, Biology, Wake Forest University
Expected graduation: May 2009
Advisor: Dr. William K. Smith
TEACHING
Certification:
6-12 Biology, State of Florida Teaching License
Lecture Positions:
Current Topics in Science Research (2004-2005)
Young Middle Magnet School of Math, Science, and Technology;
Tampa, FL
Biology (2001-2002)
Deltona High School; Deltona, FL
Integrated Science (2001-2002)
Deltona High School; Deltona, FL
7th Grade Science (2001)
DeLand Middle School; DeLand, FL
Laboratory Classes Taught:
Plant Systematics (2008); Wake Forest University
Plant Ecophysiology (2006) ; Wake Forest University
Comparative Physiology (2005-2007); Wake Forest University
General Biology I (2002-2004); Appalachian State University
General Biology II (2002-2004); Appalachian State University
General Biology I (2003); Caldwell Community College
Guest Lecturer:
Plant Physiological Ecology, Wake Forest University (2007)
Environmental Science, R.J. Reynolds High School (2008)
153
RESEARCH
Publications
Hughes NM, Neufeld HS, Burkey KO. 2005. Functional role of anthocyanins in high-light
winter leaves of the evergreen herb Galax urceolata. New Phytologist 168: 575-587.
Hughes NM, Smith WK. 2007. Attenuation of incident light in Galax urceolata
(Diapensiaceae): concerted influence of adaxial and abaxial anthocyanic layers on
photoprotection. American Journal of Botany 94: 784-790.
Hughes NM, Morley CB, and Smith WK. 2007. The coordination of anthocyanin decline
and photosynthetic maturation in developing leaves of three deciduous tree species. New
Phytologist 175: 675-685.
Hughes NM, Smith WK. 2007. Seasonal photosynthesis and anthocyanin production in ten
broadleaf evergreen species. Functional Plant Biology 34: 1072-1079.
Hughes NM, Vogelmann TC, Smith WK. 2008. Optical effects of abaxial anthocyanin on
absorption of red wavelengths by understory species: re-visiting the back-scatter hypothesis.
Journal of Experimental Botany 59: 3435–3442.
Smith WK, Hughes NM. 2008. Progress in coupling plant form and photosynthetic
function. Castanea (in press).
Hughes NM, Johnson DM, Akhalkatsi M, and Abdaladze O. 2008. Characterization of
seedling microsites at the alpine-treeline ecotone for Betula litwinowii, a deciduous treeline
species. Arctic, Alpine, and Antarctic Research (in press).
Archetti M, Döring T, Hagen S, Hughes N, Leather S, Lee D, Lev-Yadun S, Manetas Y,
Ougham H, Schaberg P, Tomas H. 2008. Adaptive explanations for autumn colours- An
interdisciplinary approach. Trends in Ecology and Evolution (accepted).
Manuscripts in Preparation
Hughes NM, Vogelmann TC, SmithWK. Attenuation of green light by abaxial
anthocyanin reduces photoinhibition under high-light in understory species.
Hughes NM, Burkey KO, Smith WK. Seasonal antioxidant and xanthophyll cycle
characteristics of broadleaf evergreen species differing in anthocyanin production during
winter.
Hughes NM, Reinhardt KS, Gierardi AR, and Smith WK. Comparative water relations
broadleaf evergreen species differing in production of anthocyanin in winter.
154
Paper Presentations
Hughes NM, Smith WK. "Winter color change: why do some evergreen species synthesize
anthocyanin in winter leaves, while others don't?" Ecological Society of America, oral
presentation (Milwaukee, WI 2008).
Hughes NM, Smith WK. "Winter color change: the adaptive role of anthocyanin pigments
in broad-leaf evergreen species". Botanical Society of America, oral presentation
(Vancouver, BC 2008).
Hughes NM, Morley CB, Smith WK. "Red coloration in developing leaves: Coordination
of anthocyanin decline and photosynthetic maturation". Botanical Society of America,
poster presentation (Vancouver, BC 2008).
Hughes NM, Smith WK, and Vogelmann TC. "The adaptive significance of red abaxial
coloration in understory plants". Association of Southeastern Biologists, oral presentation
(Spartanburg, SC 2008).
Hughes NM, Smith WK, Reinhardt KS, and Burkey KO. "Winter color change in
'evergreen' leaves: The adaptive significance of anthocyanin pigments". Association of
Southeastern Biologists, poster presentation (Spartanburg, SC 2008).
Hughes NM. "Why leaves turn red: The functional significance of anthocyanin pigments in
photosynthetic adaptation". Wake Forest University departmental seminar (2008).
Hughes NM. "Winter color change: Functional role of anthocyanins in evergreen species".
Oxford University workshop on autumnal leaf colouration (2008).
Hughes NM, Smith WK, Vogelmann TC. "Abaxial anthocyanin: contrasting adaptive
function in high light and low light species". Ecological Society of America, oral
presentation (San Jose, CA 2007).
Hughes NM, Smith WK, Vogelmann TC. "Why undersurfaces of leaves are sometimes red:
contrasting the adaptive function of abaxial anthocyanin in high light and low light species
". Botanical Society of America, oral presentation (Chicago, IL 2007).
Hughes NM, Smith WK. "When are red juvenile leaves mature enough to be green? The
coordination of anthocyanin decline and photosynthetic maturation". Association of
Southeastern Biologists, oral presentation (Columbia, SC 2007).
Hughes NM, Smith WK. "Contrasting the adaptive function of abaxial anthocyanin in
leaves adapted to high-light and low-light environments". Ecological Society of America,
oral presentation (Memphis, TN 2006).
155
Hughes NM, Smith WK. "The coordination of anthocyanin decline and photosynthetic
maturation in developing leaves of three deciduous tree species". Wake Forest University
Graduate Student Research day, poster presentation (2006).
Hughes NM. "Functional role of anthocyanin in high-light, winter leaves of the evergreen
herb, Galax urceolata". Appalachian State University departmental seminar (2004).
Hughes NM, Neufeld HS. "Ecophysiological and biochemical function of anthocyanins in
high-light winter leaves of Galax urceolata". Association of Southeastern Biologists, oral
presentation (Memphis, TN 2004).
Hughes NM. "Functional role of anthocyanin in high-light, winter leaves of the evergreen
herb, Galax urceolata". Sigma Xi, ASU chapter annual meeting invited seminar (2004).
Hughes NM. "Prevelance and frequency of temporomandibular joint disorders in musicians:
psychological and mechanical stress implications". Stetson University departmental
seminar (2001).
Hughes NM. "Prevelance and frequency of temporomandibular joint disorders in musicians:
psychological and mechanical stress implications". Stetson University Scholarship and
Performance Day (2001).
Awards
Physiological Section Award for Best Poster
-2008 Botanical Society of America meeting
Elton C. Cocke Award for Outstanding Graduate Student in Biology
-Wake Forest University (2008)
Best Student Paper Presentation In Botany
-2008 Association of Southeastern Biologists meeting
Best Student Poster In Botany
-2008 Association of Southeastern Biologists meeting
Euguene P. Odum Award for Best Student Paper Presentation in Ecology (runner-up)
-2008 Association of Southeastern Biologists meeting
Quarterman-Catherine Keever Award for best Student Poster in Ecology
-2008 Association of Southeastern Biologists meeting
Billings Award for Best Ecophysiological Paper Presentation (runner-up)
-2007 Ecological Society of America meeting meeting
156
Conant Award for Botanical Imaging (third place)
-2007 Botanical Society of America meeting
Eugene P. Odum Award for Best Student Paper in Ecology
-2004 Association of Southeastern Biologists meeting
Outstanding Graduate Research in the Sciences
-ASU chapter of the Sigma Xi Scientific Research Society (2004)
Outstanding Graduate Teaching Assistant Award
-Appalachian State University (2004)
Biology Department’s Excellence in Research Award
-Stetson University (2001)
Outstanding Senior in the Department of Biology
-Stetson University (2001)
Randolph L. Carter Award- Outstanding Secondary School Intern
-Stetson University (2001)
Runner-up for best oral presentation at the 2001 Stetson Scholarship & Performance Day
-Stetson University (2001)
Award for outstanding contribution to music by a non-major
-Stetson University School of Music (2001)
Grants and Funding
Richter Fellowship for International Research
-Wake Forest University (2008)
Student Travel Award
-Association of Southeastern Biologists (2004, 2008)
Elton C. Cocke Travel Award
-Wake Forest University, Biology Department (2006, 2007, 2008)
Vecellio Award
- Wake Forest University (2006, 2007, 2008)
Alumni Travel Award
-Wake Forest University (2006, 2007, 2008)
157
Professional Affiliations:
Association of Southeastern Biologists
Botanical Society of America
Sigma Xi Scientific Research Society
Ecological Society of America
Beta Beta Beta biological honors society
FINE ARTS
Visual Art:
A Minimalist Evening- Performance Art with Pianist Bruce Moser
-University of North Carolina, Greensboro (2008)
Wake Forest University Student Art Exhibition
-Charlotte and Philip Hanes Gallery, Wake Forest University (2008)
Artists on Liberty Gallery featured artist- Solo Show
-Artists on Liberty Gallery, Winston-Salem, NC (2008)
Science as Art Exhibition
-Chelsee’s Coffee Shop, Winston-Salem, NC (2008)
Music:
Principal Bassoon, Stetson University Orchestra (1999-2001)
Principal Bassoon, Stetson University Symphonic Band (1997-2001)
Blue Banana Quintet, Stetson University (1998-2001)
Music Teacher, Day Spring Summer Camp (2000)
158