Download Temperature-induced bleaching of corals begins with impairment of

Plant, Cell and Environment (1998) 21, 1219-l 230
Temperature-induced bleaching of corals begins with
impairment of the CO 2 fixation mechanism in zooxanthellae
R. J JONES,’ 0. HOEGH-GULDBERG,’ A. W. D. LARKUM’ & U. SCHREIBER’
‘School of Biological Sciences, The University o f Sydney, Sydney NSW 2006, Australia, and 2Julius-von-Sachs Institut fur
Biowissenschaften, Universitiit W&burg, Mittlerer Dallenbergweg 64, D-97082 Wiirzburg, Germany
ABSTRACT
INTRODUCTION
The early effects of heat stress on the photosynthesis of
symbiotic dinoflagellates (zooxanthellae) within the tissues of a reef-building coral were examined using pulseamplitude-modulated (PAM) chlorophyll fluorescence
and photorespirometry. Exposure of Stylophora pistillata
to 33 and 34 o C for 4 h resulted in (1) the development of
strong non-photochemical quenching (qN) of the chlorophyll fluorescence signal, (2) marked decreases in photosynthetic oxygen evolution, and (3) decreases in optimal
q u a n t u m y i e l d (FJF,) of photosystem II (PSII).
Quantum yield decreased to a greater extent on the illuminated surfaces of coral branches than on lower
(shaded) surfaces, and also when high irradiance intensities were combined with elevated temperature (33 “C as
opposed to 28 oC). qN collapsed in heat-stressed samples
when quenching analysis was conducted in the absence of
oxygen. Collectively, these observations are interpreted
as the initiation of photoprotective dissipation of excess
absorbed energy as heat (qN) and O,-dependent electron
flow through the Mebler-Ascorbate-Peroxidase cycle
(MAP-cycle) following the point at which the rate of
light-driven electron transport exceeds the capacity of
the Calvin cycle. A model for coral bleaching is proposed
whereby the primary site of heat damage in S. pistillata is
carboxylation within the Calvin cycle, as has been
observed during heat damage in higher plants. Damage
to PSII and a reduction in FJF, (i.e. photoinhibition)
are secondary effects following the overwhelming of photoprotective mechanisms by light. This secondary factor
increases the effect of the primary variable, temperature.
Potential restrictions of electron flow in heat-stressed
zooxanthellae are discussed with respect to Calvin cycle
enzymes and the unusual status of the dinoflagellate
Rubisco. Significant features of our model are that (1) damage to PSII is not the initial step in the sequence of heat
stress in zooxanthellae, and (2) light plays a key secondary
role in the initiation of the bleaching phenomena.
Bleaching is synonymous with the loss of pigmentation in
reef-building corals. This loss of pigmentation usually
occurs due to a decrease in the number of symbiotic
dinoflagellate algae (zooxanthellae) in the tissues of the
host (Yonge & Nicholls 1931; Hoegh-Guldberg & Smith
1989; Kleppel, Dodge & Reese 1989). It may also occur as
a result of a decrease in the concentration of photosynthetic
pigments in the zooxanthellae (Hoegh-Guldberg & Smith
1989; Kleppel et al. 1989). Coral bleaching is considered to
be a reaction to abnormal environmental conditions and has
been observed in response to variation in a wide range of
physical and chemical parameters (Yonge & Nicholls 1931;
Hoegh-Guldberg & Smith 1989; Jones 1997a). However,
the widespread bleaching of corals on individual reefs
(localized bleaching events), or the bleaching of many reefs
over large geographic areas (‘mass-bleaching’ events), has
largely been correlated with elevated sea surface temperature (Glynn 1993; Brown 1997). Bleaching has often been
observed where average daily seawater temperatures
exceed the mean summer maximum temperature by as little
as l-2 o C (Hoegh-Guldberg & Salvat 1995; HoeghGuldberg, Berkelmans & Oliver 1996; Brown, Dunne &
Chansang 1996; Jones, Berkelmans & Oliver 1997).
The mechanism that underpins this thermal sensitivity
has been the subject of considerable attention (Jokiel &
Coles 1974, 1977; Hoegh-Guldberg & Smith 1989; Glynn
& D’Croz 1990; Jokiel & Coles 1990), with many studies
focusing on how elevated temperature affects the photosynthesis of the zooxanthellae (Iglesias-Prieto, Matta &
Trench 1992; Fitt & Warner 1995; Iglesias-Prieto 1995;
Warner, Fitt & Schmidt 1996). Recent studies have used
chlorophyll fluorescence techniques to measure the ratio of
variable/maximum fluorescence (Fv/Fm), which is an indicator of PSII photochemical efficiency or the quantum
yield of photochemistry (Bjorkman & Demmig 1987;
Schreiber & Neubauer 1990; Krause & Weis 1991; Gquist,
Anderson, McCaffery & Chow 1992). In addition, chlorophyll fluorescence quenching analysis by the so-called
‘saturation-pulse technique’ has allowed for the separation
of photochemical (qP) and non-photochemical (qN) components of overall chlorophyll fluorescence quenching
(Schreiber, Schliwa & Bilger 1986). These components
have proved particularly informative about the regulatory
Key-words: Zooxanthellae; bleaching; Calvin cycle; coral;
fluorescence; Mehler reaction; Rubisco.
Correspondence: Dr Ove Hoegh-Guldberg. Fax: (02) 9351 4119;
e-mail: [email protected]
0 1998 Blackwell Science Ltd
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R. J. Jones et
al.
processes that occur within the photosynthetic apparatus of
plants, especially when these organisms are under stress
(Schreiber, Bilger & Neubauer 1994; Foyer, Lelandis &
Kunert 1994).
The application of PAM fluorometry to the study of symbiotic zooxanthellae has been provocative. Iglesias-Prieto
et al. (1992) reported a cessation in photosynthetic oxygen
evolution and loss of variable fluorescence in cultured
zooxanthellae exposed to temperatures of 34-36 “C. Fitt &
Warner (1995) and Warner et al. (1996) measured similar
effects in zooxanthellae within the tissues (in hospite) of a
number of Caribbean corals exposed to water temperatures
of 32 and 34 “C. In a more diagnostic analysis of the
effects of heat stress, Warner et al. (1996) observed that
corals with zooxanthellae which were able to develop
strong non-photochemical quenching of chlorophyll fluorescence were more temperature-tolerant (see also Fitt &
Warner 1995). Non-photochemical quenching is considered to be a photoprotective mechanism which functions to prevent the over-reduction of the photosynthetic
electron transport chain by dissipating excess absorbed
light energy in the PSII antenna system as heat
(Demmig-Adams 1990). Several components of qN have
been observed. Under most physiological conditions,
however, the largest component is referred to as ‘energydependent’ quenching. This is correlated with the
‘energization’ of the thylakoid membrane, i.e. with the
formation of a proton gradient between the stroma and
thylakoid membrane (ApH) that occurs during lightdependent electron flow (Mitchell 1961). The mechanism
by which a decrease in lumenal pH (increase in ApH)
allows de-excitation of light-generated excited states is
thought to involve the xanthophyll cycle (Demmig et al.
1987; Demmig-Adams 1990), possibly brought about by
structural changes in aggregations of light-harvesting
complexes (Horton & Ruban 1994).
Iglesias-Prieto (1995) and Warner et al. (1996) have suggested that the initial site of damage to the photosynthetic
apparatus by elevated water temperature is located within
the PSII reaction centre complex. The exact cause of damage to the reaction centre has not been determined. If analogous to photoinhibition in higher plants, however, it may
involve an increase in the reduction state of the primary
electron acceptor Q A , during acceptor side inhibition, or
the inactivation of the water-splitting system by calcium
depletion during donor side inhibition (Styring &
Jegerschold 1994). It is thought that metabolic disturbances that cause decreases in the rate of electron flow
through the transport chain to the dark-reactions of photosynthesis (Calvin cycle) carry the risk of strongly reducing
the acceptor side. The net result of this is that the potential
for photodamage at the PSI1 reaction centre increases
(Demmig-Adams 1990; Walker 1992). For example, a
decrease in the quantum yield associated with heat stress in
higher plants has been shown to occur after initial damage
to electron flow beyond photosystem I (PSI; Schreiber &
Bilger 1987). In this case, photochemical quenching
almost decreases to zero while a major part of light energy
trapped by the photosynthetic apparatus of heat-stressed
leaves is dissipated into heat. The latter is revealed by
strong non-photochemical quenching. These changes
occur before irreversible damage to PSII, and hence imply
that the earliest steps of heat damage in higher plants are
associated with limitations on electron flow through the
Calvin cycle (Weis 198 1; Schreiber & Bilger 1987; Bilger,
Schreiber & Lange 1987). The important point here is that
damage to PSI1 occurs following redox imbalance that was
caused by the initial disruption of CO,-fixation.
In the present study, modulated chlorophyll fluorescence
and photorespirometry were used to explore the effects of
heat stress on the photochemistry of the zooxanthellae
within the tissue of the coral Stylophora pistillata. The
results of this study reveal that initial damage during heat
stress occurs in the rate of the dark reactions (Calvin cycle)
of photosynthesis, preceding damage to PSII. Alterations
in assimilatory flow beyond PSI are hence the earliest step
discovered so far in the dysfunction of symbiont photosynthesis of reef-building corals by thermal stress.
MATERIALS AND METHODS
Experiments were carried out at One-Tree Island Research
Station (23”31’S, 152”08’E, Great Barrier Reef, Australia)
using the hermatypic coral Stylophora pistillata. Small
branches (50-60 mm long) were collected from individual
colonies located at 0.5-1 m depth. The pieces were
mounted using modelling clay into small plastic holders
and left at 2 m depth in One-Tree Island lagoon overnight
before use in each experiment.
Experimental design
Two experiments were carried out in which corals were
exposed to elevated water temperatures outdoors under
natural sunlight. Experiments were conducted in opentopped glass aquaria, screened from ultraviolet (IJV) light
using acrylic covers (cut off ~400 nm, Lesser & Shick
1989). Two corals were placed in each of two duplicate
aquaria at each temperature tested (experiment 1: 28, 32
and 34 “C; experiment 2: 28,30,32 and 34 “C) for a period
of 4 h. Seawater was aerated using air pumps and was
heated and recirculated using submersible pumps and
aquarium heaters. Refrigerated coolers and refrigerated
water-baths were used to control the temperatures to within
+ 0.3 “C of the desired level. Light levels (photosynthetically active radiation, PAR, 400-700 nm) in quanta (pm01
quanta mm2 ss’) were recorded with a LI-190SA quantum
sensor at 15 min intervals during experiment 1, and logged
at 10 s intervals onto a LI-COR LIlOOO data-logger during
experiment 2.
Two additional experiments were carried out in which
corals were exposed to elevated water temperatures under
artificial light (400 pm01 quanta me2 s-‘) provided by 50 W
spotlights. Corals (n = 4 at each temperature) were exposed
to 28,30,32,33 and 34 “C for 1 or 4 h (experiment 1) and
28, 30, 32, 33 and 34 “C for 4 h (experiment 2).
0 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 1219-1230
Temperature-induced bleaching of corals
Refrigerated water-baths were used to control the temperatures to within a.1 “C of the desired levels.
The interaction of light, temperature and colony branch
orientation was investigated using freshly collected tips of
S. p i s t i l l a t a . Ten colonies were orientated horizontally
within 1 L glass beakers filled with filtered seawater and
then exposed to 50,200,400,800 or 1600 pm01 quanta me2
s-i for 3 h at temperatures of 28 or 33 “C. Light intensities were produced by a combination of 50 W spotlights
and neutral density filters (50% normal density). Corals
were dark-adapted before measurement of the maximum
potential quantum yield (see below) on the upper surface
of five randomly selected corals and the lower surface of
the remaining five branches.
Measurements of chlorophyll fluorescence
Chlorophyll fluorescence was measured using two different types of pulse amplitude modulation (PAM) fluorometers (TEACHING-PAM and DIVING-PAM Walz,
Effeltrich, Germany; Schreiber, Schliwa & Bilger 1986).
Corals were placed in darkness (dark-adapted) for 20 min
prior to each measurement. They were then placed adjacent
to the fluorometer optical head and the initial fluorescence
(F,) was measured by applying a weak pulsed red light
(< 1 pmol quanta mm2 s-l). A saturating pulse (600 ms) of
actinic light (3000 prnol quanta mm2 ss’) was then applied
to a small section of the coral surface in order to measure
maximum fluorescence (F,,,). Variable fluorescence (F,)
was calculated as F,,, - F,, and maximum potential quantum
yield as FJF,,,. Quenching analysis of zooxanthellae
within the corals was explored using the TEACHINGPAM. This instrument was used to measure the photochemical (qP) and non-photochemical (qN) quenching
components during induction curve analysis. In this case, a
series of saturating flashes at 20 s intervals were applied to
measure F,,,’ immediately after dark adaptation and during
exposure to set PAR irradiances. Photochemical quenching, qP = (F,,,’ - F)/(F,,,’ - F,), non-photochemical quenching, qN = (F,,, - F,‘)/(F,,, - F,,), and yield (AFIF,‘) were
calculated according to Schreiber et al. (1986) using the
nomenclature described in van Kooten & Snel(l990).
Measurement of oxygen flux
The effect of elevated water temperature on the photosynthesis and respiration of experimental corals was measured
using a laboratory-based ‘photorespirometer’ comprising
four separate 90 mL water-jacketed acrylic chambers. Each
chamber had a false bottom enclosing a stir bar powered by
a magnetic stirrer. Actinic light was provided by 50 W
spotlights, illuminating each chamber from opposite sides.
Respiratory O2 consumption and photosynthetic O2 production were measured using Clark-type electrodes
(Strathkelvin Instruments, Glasgow) inserted into the
chamber tops. Sensors were connected via oxygen-polarizing units to an analogue to digital converter (ADC-1,
Remote Measurement Systems, Seattle, USA) which was
0 1998 Blackwell Scicncc Ltd, Plant, Cell and Environment, 21, 1219-1230
1221
controlled by data acquisition software (DATACAN IV,
Sable Systems, Los Angeles) run by an IBM-compatible
laptop computer. Oxygen concentrations were measured
every 6 s from an average of 2 consecutive voltage readings from each sensor. The voltage of the sensors was calibrated using air-saturated seawater at the incubation
temperature and salinity and oxygen purged (nitrogen-bubbled) seawater. Oxygen concentrations at saturation were
computed for each experimental temperature and salinity
(Temptabl program 1986, Oxford University Press).
Respiratory and photosynthetic rates were measured by
monitoring changes in dissolved 0, concentrations over
10 -12-min periods. Oxygen consumption in the dark was
measured before and after exposure to saturating light
(750 pm01 quanta mm2 ss’, see Fig. 5). The average of the
two dark measurements was used to calculate gross photosynthesis from the net photosynthetic rates. All measurements of respiration and photosynthesis were taken at the
temperatures at which the corals were previously incubated (i.e. 28,30,32 or 34 “C). Where chlorophyll fluorescence parameters were measured after photorespirometry
(see ‘Results’), corals were dark-adapted at the incubation
temperatures (28,30,32 or 34 “C) for 20 min prior to measurement of maximum potential quantum yield. Quenching
analysis was conducted at ambient temperature (28 “C ).
Photosynthetic rate versus irradiance (P-r) curves were
measured for six corals immediately after collection.
Corals were exposed to darkness or to seven different irradiance levels (~16-500 bmol quanta mm2 s ‘) obtained by
inserting a number of neutral density filters between the
lights and incubation chambers housing the corals.
Biomass analysis
Corals were frozen for measurement of the number of
zooxanthellae in the tissue and concentration of chlorophyll immediately after experimentation. A subset of corals
were also frozen immediately after the collection of the
branch tips to measure the population density of zooxanthellae and concentration of chlorophyll a (Chl a) in the
parent colonies (PC). From these parameters, it could be
determined whether the handling and preparation procedures caused any significant loss of zooxanthellae.
Tissues were stripped from the skeletons with a jet of
recirculated 0.45 ,um membrane-filtered seawater using a
WaterPik TM (Johannes & Wiebe 1970). The slurry produced from the tissue-stripping process was homogenized
in a blender for 30 s and the volume of the homogenate
recorded. The number of zooxanthellae in 10 mL aliquots
of the homogenate was measured using a hemocytometer
(8 replicate counts). The total number of zooxanthellae per
coral was measured after correcting for the volume of the
homogenate. Zooxanthellae density was calculated as
number per unit surface area. Coral surface area was measured using the paraffin wax technique (Stimson & Kinzie
1991). For Chl a analyses, two 10-20 mL subsamples of
the homogenate were filtered through Whatman GF/C filters which were then homogenized for 30 s using a tissue
1222
R. J. Jones
et al.
homogeniser. Chl a was extracted with 10 mL of acetone
for 24 h in a freezer. Extracts were centrifuged to remove
filter fibres from suspension and the supematants read on a
spectrophotometer using the equations of Jeffrey &
Humphrey (1975) to calculate the concentration of Chl a.
RESULTS
Observations of the natural bleaching of corals at
One-Tree Island
There have been no documented bleaching events at OneTree Island; however, widespread bleaching of corals was
noted independently by two of the authors (Jones &
Hoegh-Guldberg) during early February 1996 and
(recently) by M. Waugh in April 1998 (M. Waugh, manager, One-Tree Island Research Station, personal communication). In situ water temperatures have been recorded at
Heron Island, 20 km north of One-Tree Island, since
December 1995 as part of the Great Barrier Reef Marine
Parks Authority (GBRMPA) Sea Temperature Monitoring
Program (GBRMPA, Research and Monitoring Section).
Sea water temperature recorded at 0.5 h intervals on the
reef flat (1 m depth) by platinum thermocouples (accuracy
kO.1 “C) were obtained from the GBRMPA for the period
December 1995 to May 1996. The in situ recordings indicate a period of 6 d between 28 January and 3 February
1996 in which the maximum daily seawater temperature
exceeded 32 “C. The highest absolute water temperature
measured over the period was 34.0 “C (4.30 pm 30 January
1996, Fig. 1), which was also the highest maximum water
temperature recorded over the summer (December
1995-March 1996). On that day, water temperatures
exceeded 33 “C for ~3 h and 32 “C for ~4 h (Fig. 1 inset).
There was no discoloration (bleaching) of the corals
following exposure to elevated water temperatures under
UV-screened solar radiation. However, zooxanthellae densities in corals exposed to 34 “C for 4 h were = 60% of the
densities measured in corals exposed to ambient water
temperatures (28 “C), or branch tips sampled from the parent colonies (Fig. 2). Chl a concentrations per zooxanthellae did not differ between treatments in either experiment
(data not shown). There were no significant differences
between the algal densities of control corals (28 “C) and
parent colonies, suggesting that the preparation and manipulative procedures had no measurable effect on the loss of
zooxanthellae from the test corals.
The first and second thermal experiments were conducted
during sunny days with intermittent cloud cover. Light levels during the first experiment were recorded every 15 min,
and minimum and maximum values of 500 and 1500 pm01
quanta mm2 s& were obtained. Light levels during experiment 2 were logged at 10 s intervals (see Fig. 2, inset) and
varied between 390 and 1540 pmol quanta mm2 s-‘.
The maximum potential quantum yield (F,/F,,,) of the
28 “C-treated control corals was similar to levels measured
in freshly collected corals (range 0.56-0.64, n = 10). ‘FJF,,,
in the 34 “C-treated corals was significantly different from
that of control corals in both experiments (Fig. 3, ANOVA
P < 0.05), corresponding to a = 40% (experiment 1) and
= 21% (experiment 2) reduction in FJF,,,. In experiment 1,
FJF,,, in the 32 “C-treated corals was also significantly
different from that of control corals (P < 0.05).
Representative fluorescence traces of control (28 “C)
and 34 “C-treated corals from the experiments conducted
under UV-screened solar radiation (experiment 1 above)
are shown in Fig. 4. Fm‘, qN and yield (AFIF,‘) were measured by the saturation-pulse technique using the TEACHING-PAM chlorophyll fluorometer. Fluorescence yield
was monitored by applying a weak modulated measuring
beam. Shortly before the recording, the F, level was measured and a saturation pulse was applied to assess F,,, and
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Figure 1. Seawater temperature (“C) at Heron Island (20 km north of One-Tree Island) recorded every 0.5 h at 1 m depth on the reef flat. Main
figure: water temperatures between 1 January 1996 and 4 March 1996. Inset: water temperatures between 30 January and 1 February 1996.
0 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 1219-1230
Temperature-induced bleaching of corals 1223
= 9 #*
+ Experiment 1
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30
32
34
36
Temperature (“C)
Figure 2. Mean zooxanthellae cm* in Stylophora pistillata
following exposure to water temperatures of 28,30,32 and 34 “C
for 4 h under UV-screened solar radiation. Data are means of six
replicates (experiment 1) or four replicates (experiment 2) * 95%
confidence intervals (CI). Data points have been shifted slightly
along the x-axis for clarity. #P = 0.06; * P <# 0.05. Inset: PAR during
experiment 2; the arrow indicates the duration of the experiment.
the maximum potential yield, FJF,,,, of the dark-adapted
sample (see Fig. 3). When the fluorescence yield had
relaxed almost back to the original F, level, actinic illumination was turned on (at ~ 20 s in Fig. 4) and the lightinduced changes of fluorescence yield were measured.
Induction of a normal ‘Kautsky effect’ (Kautsky & Hirsch
1931) was evident in the control explant (Fig. 4a).
Fluorescence yield first rapidly rose to a primary and then
to a secondary peak before it slowly declined to a steadystate level. Saturating pulses were applied every 20 s in
order to measure F,‘, which is reduced with respect to F,,,
by qN. The values of effective quantum yield, AFIF,‘, and
of qN are also displayed. In the control sample, during the
first minute of illumination, AFIF,’ was almost constant,
whereas qN increased. The following rise of AFIF,’ was
correlated with a decrease of qN. These features are well
known from previous work on higher plant leaves (e.g.
Schreiber & Bilger 1987). The initial rise of qN to a high
level reflects the build-up of a substantial ApH, as Calvin
cycle enzymes have yet to be activated by light. As a result
of this, ATP that would normally be consumed by the dark
reactions accumulates. Non-assimilatory electron flow
must be responsible for initial ApH formation. As Calvin
cycle activation occurs, ATP is consumed, the ApH is utilized and there is a corresponding decrease of qN, accompanied by an increase of AFIF,,,‘.
In the 34 oC-treated sample (Fig. 4b), the non-photochemical quenching of fluorescence yield that is induced
by light is considerably enhanced, such that F,’ almost
reaches the original F, level. At the same time the effective
quantum yield, AFIF,‘, is decreased with respect to the
control. Notably, qN does not relax during illumination and
after an initial rapid rise further increases during a second
slower phase. Hence, there is no induction of Calvin cycle
0 1998 Blackwell Science Ltd. Plant, Cell andEnvironment, 21, 1219-1230
activity by analogy to previous observations in the study
of heat-stressed leaves (Schreiber & Bilger 1987; Bilger
et al. 1987). It should be noted, however, that a fairly
high maximum effective quantum yield is maintained even
in the 34 “C sample, which reflects non-assimilatory,
energizing electron flow.
In Fig. 4c the nature of this non-assimilatory electron flow
is revealed by an analysis conducted with a 34 “C-treated
sample under saturated nitrogen (low oxygen). It is revealed
that the build-up of ApH, which is normally reflected by a
rapid rise in qN (see Fig. 4b), is dramatically slowed down.
At the same time, the maximum effective quantum yield,
AFIF,‘, is substantially suppressed. This may be taken as
evidence for a decisive role of oxygen-dependent electron
flow in thylakoid membrane energization and for the excessive non-photochemical quenching observed in the heattreated sample. It cannot be ruled out that inhibition of
respiration in both the host and endosymbiont cells may also
feed back via ATP/ADP and NAD(P)H/NAD(P) levels to
photosynthetic electron transport (see ‘Discussion’).
Both of the experiments conducted under UV-screened
solar radiation were carried out on cloudy days in which
PAR levels varied considerably (see Fig. 2, inset). To
examine further the response of zooxanthellae from S. pistillata to elevated water temperature, we repeated the
experiments, but this time under controlled lighting conditions provided by 50 W spotlights. The photosynthetic
response of zooxanthellae from S. pistillata as a function of
irradiance (Fig. 5) was initially examined to ensure that
experimental light levels used only just saturated the photosynthetic processes, and hence were not too stressful.
A saturating light level of 400 pm01 quanta mm2 s-l was
chosen from the P-I curve (Fig. 5), and corals were
0.8
+-Experiment 1
+ Experiment 2
0.7 0.6 I
LLE
. 0.5 LL5
0.4 0.3
0.2
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30
32
Temperature (“C)
34
Figure 3. FJF, of corals exposed to water temperatures of 28,30,
32 or 34 “C for 4 h under UV-screened sunlight. Data are means of
six replicates (experiment 1) or four replicates (experiment
2) f 95% CI. Data points have been shifted slightly along the n-axis
for clarity. * Significant, P < 0.05. Corals were dark-adapted for
20 min and fluorescence parameters measured at the same
temperature as that to which the corals were previously exposed.
R. J. Jones et al.
FJF,,, of corals exposed to 34 “C for 1 or 4 h was significantly different from that of the 28”C-treated control
corals (Fig. 7), corresponding to a ~ 20% reduction of
F,/F,. A similar reduction of F,/F, was measured in
corals exposed to 34 “C during the repeated experiment
(Fig. 7 inset; see also above).
As shown in Fig. 8, the decrease in F,/F,,, observed in
heat-stressed corals (Fig. 7) was mainly caused by a
decrease in F,,,, whereas there was only a small increase in
the F, level.
Quenching analysis (conducted on corals from experiment 2 only) revealed the development of strong qN in the
34 “C-treated samples (P < 0.05, Fig. 9), similar to that
observed in corals exposed to 34 “C under UV-screened
sunlight (Fig. 4b). Photochemical quenching (qP) was not
significantly different between treatments (Fig. 9).
Maximum potential quantum yield of zooxanthellae from
the upper (illuminated) and lower (shaded) surfaces of S.
pistillata was measured in corals exposed to five different
irradiance levels for 3 h at 28 or 33 “C. FJFm on the illuminated surface of the corals in the 33 “C treatment was consistently lower than in the 28oC-treated corals (Fig. 10).
Significant differences between the two temperature treatments were recorded at each irradiance level (ANOVA
P > 0.05). In the 28oC-treated samples, FJF,,, on the upper
surfaces of corals was significantly different from that on the
shaded surface following exposure to 800 and 1600 ~01
quanta me2 s-l (Fig. 10). This corresponded to a 13 and 26%
reduction in maximum potential quantum yield (Fig. lOa).
In the 33oC-treated corals F,/F,,, on the upper surfaces
0.5 -
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0
e
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0 50 100
150
200
250
300
Time (s)
Figure 4. Dark-light induction curves with saturation-pulse
quenching analysis for (a) control colony maintained at 28 “C (b)
colony exposed to 34 “C for 4 h, and (c) 34 “C-treated sample kept
in an atmosphere of nitrogen for 5 min. Measurements were made
with the TEACHING-PAM chlorophyll fluorometer. Corals were
dark-adapted for 20 min at the same temperature as that to which
the corals were previously exposed. Quenching analysis was
conducted at ambient temperature. The displayed recordings were
preceded by F, - F,,, determinations involving the application of
saturation pulses 30 s before the start of recording at time zero. The
characteristic levels of fluorescence yield and fluorescence
parameters are denoted in (c) with the following definitions: F,.
minimum fluorescence yield of dark-adapted sample; F,,,, maximum
fluorescence. yield of dark-adapted sample; F,,,‘, maximum
fluorescence yield of illuminated sample; F, time-dependent
fluorescence yield; qN, coefficient of non-photochemical
quenching; AFIF,‘, effective quantum yield of PSII. See text for
further explanation.
1
exposed to temperatures in the range 28-34 “C for 1 and
4 h. No significant differences were detected in zooxanthellae per area or Chl a per zooxanthellae, between the
control corals (28 “C) or parent colonies (PC controls) and
any of the temperature treatments (data not shown).
In S. pistillata exposed to 34 “C for 1 or 4 h, gross photosynthetic oxygen evolution was significantly different from
that of control (28 “C-treated) corals (ANOVA, Fig. 6). This
experiment was repeated using new corals that were
mounted and prepared as described for the first experiment.
Again, gross photosynthesis of the 34 “C-treated samples
was dramatically reduced, corresponding to < 10% of the
rate measured in the control corals (Fig. 6, inset).
.o
,
4
100
200
PAR (umol
300
400
quanta mm2 s’)
500
Figure 5. Gross photosynthesis urn01 O2 zooxanthella? h-l) versus
imadiance @noI quanta m? s?) curve. for Sfylophorapistillata used
in tbe study. The curve fit is a hyperbolic tangent function (see Chalker
1981). Data are means of six replicates * 95% CI.
0 1998 Blackwell Science Ltd. Plant, Cell and Environment, 21, 1219-1230
Temperature-induced bleaching of corals
1.3
-k
‘i
i
The impact of heat stress on the photosynthesis
of zooxanthellae
Exposure time: -c- 1 h -0-4 h
T
1.0
We observed three characteristics of zooxanthellae from
heat-stressed S. pistillata using saturation-pulse PAMfluorometry and photorespirometry: (1) an increase in nonphotochemical quenching of chlorophyll fluorescence
which is not accompanied by any significant decrease of
photochemical quenching, (2) a decrease in the quantum
yield of PSII, and (3) a decrease in photosynthetic oxygen
evolution. These characteristics were repeatable and
observed in experiments conducted under both UVscreened sunlight and artificial light (see e.g. Figs 6 & 7).
The decrease in the maximum potential quantum yield
of PSII and the rate of gross photosynthesis observed in
heat-stressed S. pistillata has been reported in heatstressed zooxanthellae in culture (Iglesias-Prieto et al.
1992; Iglesias-Prieto 1995) and in the tissues of the corals
:
0”
B
g 0.8
‘L
=.
.$
2
II
Experiment 2
0.5
=.
d
e
t 0.3
a
E
0.0
27
28
29
30
31
32
1225
33
34
35
Temperature (“C)
Figure 6. Gross photosynthesis @no1 Oz zooxanthella? h-l) of
zooxanthellae in Sfylophora pistilkm exposed to water temperatures
of 28,30,32,33 or 34 “C for 1 or4 h (experiment l), or4 h
(experiment 2, inset), under artificial light (4OO~mol quanta mm2
s-l). Data are means of four replicates f 95% CI. Data points have
been shifted slightly along the x-axis for clarity. * P < 0.05.
Measurements of photosynthesis and respiration were taken at the
same temperature in which the corals were previously incubated.
was also significantly different from that on the lower
surfaces at the same irradiances, but also at 200 pmol
quanta me2 s-’ (Fig. 10b). Curiously, there was no difference in FJF,,, between the upper and lower surfaces of
corals exposed to 400 pmol quanta me2 s-‘. It is not clear
why a similar light-related reduction of dark-adapted
yield was not observed, given the clear effects at the
200 pmol quanta mm* SK’ irradiance level (Fig. 10b). At
the highest irradiance tested (1600 pmol quanta mm2 s-l),
FJF,,, on the upper surface (0.3 f 0.1 n = 5) was only
53% of the value on the shaded parts of the coral. Similar
marked decreases in yield were observed during the
experiments conducted under UV-screened solar radiation, in which corals were exposed to maximum irradiances of 1500 pm01 quanta mm2 SK’ at 34 “C.
Montastrea annularis, M. cavernosa, Agaricia lamarcki,
A. agaricites and Siderastrea radians (Fitt & Warner
1995; Warner et al. 1996). High levels of non-photo-
chemical quenching have also been reported by Warner
et al. (1996) in heat-stressed corals (M. annuluris and S.
radians). Non-photochemical quenching is considered to
reflect a mechanism for photoprotection which is
designed to prevent the over-reduction of the photosynthetic electron transport chain by dissipation of excess
absorbed light energy in the PSII antenna system as heat
(Demmig-Adams 1990). Non-photochemical quenching
occurs when the rate of light-driven electron transport
exceeds the rate of ADP/Pi re-cycling by the dark reac-
Exposure time: + 1 h -0-4 h
0.70
0.60
E
+ 0.50
LLZ
Experiment 2
0.7
0.4
DISCUSSION
The events preceding the expulsion of zooxanthellae from
the tissues of corals that have been exposed to higher than
normal temperatures are complex. However, a common
theme emerges from recent investigations of photosynthetic function. These studies have shown that PSII is damaged early in heat stress. Our results support these
conclusions. We suggest, however, that damage to PSII in
heat-stressed zooxanthellae is a secondary effect that follows damage to assimilatory electron flow, possibly to the
first steps of the Calvin cycle.
0 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 1219-1230
0.3
0.30
/___._I
$:Z +Y--+.+
0.40
27
t
27
28
29
29
31
30
33
31
35
32
33
34
35
Temperature (“C)
Figure 7. FJF,,, of zooxanthellae in Stylophora pistillata
exposed to water temperatures of 28, 30,32,33 or 34 “C for 1 or
4 h (experiment l), or 4 h (experiment 2, inset), under artificial
light (400 ,umol quanta mm2 s-l). Data are means of four replicates
f 95% CL Data points have been shifted slightly along the x-axis
for clarity. * P < 0.05. Corals were dark-adapted for 20 min and
fluorescence parameters measured at the same temperature as that
at which the corals were previously incubated.
1226
R. J. Jones et al.
B 1100
5
a,
.>
%
3
0.7
0.6
0.5 <3
0.4 ,+
1000
900
.L
8
800
5
::
700
s!
0
2
600
I
500
&
400
7
m
300
+
200
FL,” +
0.1
0.0
27
28
29
30
31
32
33
34
35
36
37
Temperature (“C)
Figure 6. F,, F, and FJF,,, in Stylophora pistillata exposed to
water temperatures in the range 28-36 “C for 1 h under a saturating
light intensity (400 ~mmol quanta IT? s-l). Data are means of four
replicates f 95% CI
1 .o
.e 0.6
s
5
s 0.6
z,
b
g 0.4
0.2
0.0
r
I
qp:n
26
W q
30
32
Temperature (“C)
34
-I
Figure 9. Non-photochemical quenching (qN) and
photochemical quenching (qP) in zooxanthellae from Stylophora
oxygen-consuming pathways such as the MehlerAscorbate-Peroxidase (MAP) cycle and photorespiration
can reduce photodamage to the photosynthetic apparatus
(Schreiber & Neubauer 1990; Osmond & Grace 1995;
Biehler & Fock 1996; Cheeseman et al. 1997; Park et al.
1996; Polle 1996). In the MAP cycle, oxygen and H 2 O 2 (via
monodehydroascorbate) act as electron acceptors for PSI
(Mehler 1951; Asada & Badger 1984; Asada & Takahashi
1987; Miyake & Asada 1992; Schreiber er al. 1995). The
MAP cycle serves two functions, first to consume excess
electrons, thereby preventing accumulation of light-generated reductant, and secondly to establish a pH gradient
which induces an increase of energy dissipation into heat, as
reflected by non-photochemical fluorescence quenching
(Schreiber & Neubauer 1990). Our observation of the collapse of qN in heat-stressed zooxanthellae during quenching
analysis under saturated nitrogen is consistent with previous
findings that O2-dependent electron flow in the MAP cycle
is mainly responsible for the energization of the transthylakoid membrane when the rate of light-driven electron
transport exceeds the rate of enzymatic reactions of the
Calvin cycle (Schreiber & Neubauer 1990; Neubauer &
Yamamoto 1992; Schreiber, Bilger & Neubauer 1994). In
addition, it should be noted that removal of molecular oxygen is likely to suppress respiration in both host and
endosymbiont cells. This could lead to a drop in ATP/ADP
and an increase in NAD(P)H/NAD(P), both of which could
have an effect on photosynthetic electron transport. The drop
0.6
E
s
L’ 0 . 4
0.2
0.0
50
200
400
800
1600
200
400
800
1600
pisfilkzta
exposed to water temperatures of 28, 30,32 or 34 “C for
4 h under artificial light (400 @ml quanta mm* s-l). Data are means
of four replicates f 95% CI. * P < 0.05. Corals were dark-adapted
for 20 min at the same temperature treatment as that to which the
corals were previously exposed. Quenching analysis was
conducted under ambient temperature.
tions of photosynthesis (Schreiber & Bilger 1987;
Schreiber & Neubauer 1990; see also Figs 4 & 8).
It is particularly interesting that non-photochemical
quenching in heat-stressed zooxanthellae collapsed when
the analysis was conducted in a saturated nitrogen atmosphere (i.e. in the absence of oxygen; Fig. 4c). This observation implies that non-photochemical quenching in
heat-stressed corals results partially from O2-dependent
electron flow. There has been a growing realization in recent
years that the diversion of electrons into non-assimilatory,
50
PAR (pmol quanta rn-> s-‘)
Figure 10. A comparison of FJF, on upper (illuminated) and
lower (shaded) surfaces of Stylophora pistillata exposed to PAR of
50,200,400,800 and 1600 ~mmol quanta m-* s-’ for 3 h at 28 “C (a)
or 33 “C (b). Data are means of five replicates * 95% CI. *P < 0.05.
Corals were dark-adapted for 20 min and fluorescence parameters
measured at the same temperature as that to which the corals were
previously exposed.
0 1998 Blackwell Science Ltd, Plant, Cell and Environment, 21, 1219-1230
Temperature-induced bleaching of corals
in ATP/ADP may be related to the observed decrease in
energy-dependent quenching, and the increase in
NAD(P)H/NAD(P) could explain part of the apparent lack
of electron acceptors after oxygen removal.
The first steps in the onset of photosynthetic
dysfunction in corals
Iglesias-Prieto (1995) and Warner et al. (1996) suggest,
from similar evidence to that reported here, that the initial
site of action of elevated water temperature is on the oxygen evolving complex and the reaction centre of PSII, possibly at the site of the Dl protein (Warner et al. 1996). Our
results also indicate damage to PSI1 in heat-stressed corals,
shown by marked decrease in maximum potential quantum
yield at the higher temperatures used during the present
study (Figs 3 & 7). However, we suggest that this is a secondary effect, and a consequence of limitations in assimilatory electron flow. We propose that, during heat stress in
zooxanthellae of S. pistillata, electron flow through to
NADP reductase is blocked. This causes (1) an increased
electron flow to the MAP cycle, thereby maintaining some
electron flow, and (2) a rate of charge separation in PSII
that exceeds, under high light, the capacity of electron flow
of the MAP cycle. The resulting increase in ApH across the
thylakoid membrane initiates photoprotective dissipation
of excess absorbed light energy as heat which is reflected
by energy-dependent, non-photochemical quenching
(Figs 4 & 8). Thus, we suggest that the maintenance of a
sustained non-photochemical quenching of the fluorescence signal is supported by O2-dependent electron flow in
the MAP cycle. There is, however, only a finite capacity
for photoprotection, beyond which the electron transport
chain becomes over-reduced and significant damage to
PSI1 occurs. An important feature of this model for temperature stress is that damage to PSII is a secondary effect,
which occurs after a reduction to dark electron flow and the
advent of redox imbalance. If PSI1 were the initial site of
action during temperature stress, then it would seem
unlikely that reduced rates of electron flow could support
the ApH and hence the high levels of non-photochemical
quenching observed in this study. Furthermore, it should be
considered that the observed decrease of net oxygen evolution, which was accompanied by stimulated qN, was not
paralleled by any significant decrease of qP. This strongly
argues for a type of electron flow which does not lead to
net O2 evolution, but which still involves functional PSII.
All the criteria for such types of electron flow are fulfilled
by the MAP cycle.
A significant aspect of this model for bleaching is a
mechanistic explanation of the interaction between light
and temperature which has been observed in laboratory
experiments (Hoegh-Guldberg 1989; Jokiel & Coles 1990)
and during natural bleaching events. In the latter case, the
preferential paling of corals on their upper sunlightexposed surfaces (‘shade effect’) is a common observation
(Glynn 1983; Harriott 1985; for a review see Williams &
Bunkley-Williams 1990). We have shown that the effect of
0 1998 Blackwell Science Ltd. Plant, Cell andEnvironment, 21,1219-1230
1227
temperature is much greater on the upper, more directly
illuminated surfaces of heat-stressed S. pistillata which is
consistent with the observed patterns of coral discolouration during bleaching events. More recently, bleaching on
the upper sunlight-exposed surfaces of corals has been proposed to be due to the presence of a light/temperature-sensitive clade of zooxanthellae on the upper surface (Rowan
et al. 1997). Zooxanthellae from S. pistillata belong to
clade C (cf Rowan & Powers 1991), as identified by
restriction-fragment length polymorphisms (RFLPs) in
genes encoding small ribosomal RNA (srRNA; W. Loh &
0. Hoegh-Guldberg, University of Sydney, unpublished
results). There is no evidence of the complex multiclade
communities in S. pistillata from One-Tree Island as
reported by Rowan et al. (1997) for M. annularis. While
the differential susceptibility of certain clades may explain
bleaching patterns in some corals this does not explain why
mono-cladal corals like S. pistillata bleach on the upper
surfaces first (Hoegh-Guldberg & Smith 1989).
It is not directly obvious from our results at which site
heating induces a limitation in electron transport through to
the dark reactions of photosynthesis. Previous work on
leaves of higher plants has found that Calvin cycle
enzymes are sensitive to heat stress, preceding deactivation
of water splitting in PSII by ~ 5 “C (Weis 1981; Kobza &
Edwards 1987). The same conclusion was drawn indirectly
from measurements of fluorescence (Schreiber & Bilger
1987) as well as parallel measurements of fluorescence and
CO,-dependent oxygen evolution (Bilger et al. 1987). The
phenomenological ‘fingerprint’ of Calvin cycle inhibition
is a moderate drop in electron transport rate concurrent
with a strong enhancement of energy-dependent non-photochemical quenching. The results of our study parallels
these results; this is clear from comparing the similarity of
our data (Fig. 4a,b) with data for heat-stressed higher
plants (see fig. 11 of Schreiber & Bilger 1987).
Little is known of the thermal stability of Calvin cycle
enzymes of zooxanthellae; however, the recent discovery
of a very unusual and extremely unstable type of Ribulose1, 5bisphosphate carboxylase/oxygenase (Rubisco) in
zooxanthellae is particularly interesting. Rubisco from
zooxanthellae is similar to bacterial form II Rubisco
(Whitney, Shaw & Yellowlees 1995; Rowan et al. 1996);
all other known eukaryote Rubiscos are form I enzymes.
The cause of instability of the Rubisco from zooxanthellae
is currently unknown (Bush & Sweeney 1972; Whitney &
Yellowlees 1995). Recently, Rubisco activase, a soluble
protein found in the chloroplast and which regulates the
activity of Rubisco in vivo, has been implicated as the site
of temperature sensitivity in heat-stressed higher plants
(Crafts-Brandner, van de Loo & Salvucci 1997; Feller,
Crafts-Brandner, & Salvucci 1998). This represents an
extremely interesting area of future research.
Yonge & Nicholls (193 1) suggest that the dissociation of
the coral symbiosis during periods of elevated water temperature could be the result of CO 2 starvation. Both carboxylation and oxygenation of ribulose-1, 5-bisphosphate
are catalysed by Rubisco; since oxygenation competes
1228
R. J. Jones et al.
with carboxylation, the rate of carbon fixation depends on
the relative concentration of these gases. It is particularly
interesting that form II Rubiscos have a much lower specificity for CO, versus O2 (Jordan & Ogren 1981). Recently,
Whitney & Andrews (1998) revealed that the C02/02
specificity of Rubisco from the dinoflagellate Amphidinium
carterae was approximately twice as great as that of other
homomeric Rubiscos but unlikely to be sufficient to support
dinoflagellate photosynthesis without assistance from an
inorganic-carbon-concentrating mechanism (CCM). The
problem of keeping the equilibrium between carboxylase
and oxygenase activity of Rubisco in favour of carbon fixation is further compounded by the typically hyperoxic environment of the cnidarian host cell (Dykens & Shick 1982),
and potential limitation of CO 2 delivery when photosynthetic rates are high (Muscatine, Porter & Kaplan 1989;
Weis, Smith & Muscatine 1989; Lesser et al. 1994). If electron flow to the Calvin cycle is limited by CO 2 delivery
and/or the carboxylation activity of Rubisco (i.e. sink
limitation), then zooxanthellae run the risk of over-reduction of the electron transport chain at high photosynthetic
rates and irradiances as observed in our studies. These features of the dinoflagellate photosynthetic apparatus highlight the extremely sensitive (vulnerable) relationship
between the photochemical transduction of light and the
efficient processing of electrons by the dark reactions.
Coral bleaching
The loss of zooxanthellae observed in the present study in
heat-stressed corals is a well-known response (Jokiel &
Coles 1974, 1977; Hoegh-Guldberg & Smith 1989).
Equally, unusually high sea temperatures are the best
explanation for periodic mass bleaching events that have
been documented for tropical seas since 1980 (Glynn
1993; Hoegh-Guldberg & Salvat 1995; Brown 1997). Loss
of zooxanthellae occurred in the present study without any
decrease in algal chlorophyll concentration. A similar
effect has been observed in heat-treated corals in laboratory manipulations (Hoegh-Guldberg & Smith 1989; Fitt &
Warner 1995), and in the initial stages of natural bleaching
events in the Great Barrier Reef region (Lizard Island,
Hoegh-Guldberg & Smith 1989; Magnetic Island, Jones
1997b). Loss of algal chlorophyll may consequently be a
transitory or secondary effect resulting from long-term
exposure to elevated water temperature and high light
(Hoegh-Guldberg & Smith 1989; Jones 1997b).
Interestingly, a significant loss of zooxanthellae from S.
pistillata was observed in the experiments conducted in
sunlight (Fig. 2), but not in experiments conducted under
artificial light. The reason for this outcome is not clear but
it may relate to the different spectral qualities of the two
light sources (see Fitt &Warner 1995; Lesser 1997). In this
study we have examined the early effects of heat stress on
photosynthesis of zooxanthellae from a common reefcoral. We observed photoprotective mechanisms such as
non-photochemical quenching and possibly the MAP cycle
and photorespiration acting together to reduce photon
damage to the photosynthetic apparatus. Nevertheless,
marked decreases in photosynthetic oxygen evolution and
PSII photochemical efficiency occurred during exposure of
corals to temperatures observed in coral bleaching events.
We suggest a new model for coral bleaching in which
temperature stress begins with an impairment of dark
metabolism of the zooxanthellae (sink limitation), ultimately resulting in photon damage to PSII or the carbon
concentrating mechanism. Potential causes of this are the
effect of temperature on Calvin cycle enzymes and the
unusual status of the zooxanthellar Rubisco. Significant features of this model are that (1) damage to PSII is not the initial step in the sequence of heat stress in zooxanthellae, and
(2) light plays a key secondary influence.
ACKNOWLEDGMENTS
We thank the Great Barrier Reef Marine Parks Authority
for supplying the seawater temperature data for the Heron
Island reef flat. This work was supported by grants from
the Australian Research Council and Great Barrier Reef
Marine Park Authority to O.H.G.
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