Light-induced production of tuberculae on coral exposed
surface, endosymbiont types and their photosynthetic
responses in Montipora capitata
Ranjeet Bhagooli
Department of Biosciences, Faculty of Science, University of Mauritius,
Reduit, Mauritius
E-mail: r.bhagooli@uom.ac.mu
Abstract
Light has a pivotal role in sustaining reef-building corals, as these marine organisms exist
in symbiosis with unicellular photosynthetic dinoflagellates. The present study attempts
to examine the effect of light on the coral surface morphology and on the type of
symbiotic dinoflagellate harbored in the plate form of the coral Montipora capitata using
field data. Light measurements in exposed environments were approximately thirty fold
higher than those in the shaded areas. The plate forms of M. capitata found in exposed
light environment showed significantly higher number of tuberculae compared to those in
the shaded environment. Denaturing gradient gel electrophoresis (DGGE) analysis of the
internal transcribed spacer 2 (ITS2) region revealed one ITS2 type of symbiotic
dinoflagellate, C17, irrespective of the light regime from which the colonies were
collected. However, chlorophyll fluorescence measurements indicated that the corals
without tuberculae exhibited earlier and more pronounced photoinhibition. These results
suggest that although the light environment did not determine the type of endosymbiotic
dinoflagellate in M. capitata, it could possibly induce formation of tuberculae on the
surface skeleton as protoprotection. Reciprocal transplantation experiments confirmed the
induction of tuberculae formation in M. capitata by high light exposures.
Introduction
Since Triassic time, a period spanning over 200 million years, the coral animal and the
dinoflagellate algae, commonly known as zooxanthellae, have formed a symbiotic
relationship. All reef-building corals harbor zooxanthellae within their gastrodermal
tissues, including the connecting sheet, tentacles or mesenteries (Titlyanov et al. 1996).
They are photosynthetic, endosymbiotic, unicellular dinoflagellates representing
Symbiodinium spp. (Trench and Blank 1987). The zooxanthellae increase the fitness of
their coral host by enhancing calcification, mediating elemental flux and providing
photosynthetically fixed carbon (Muscatine and Cernichiari 1969). In return the
zooxanthellae benefit from the otherwise limited inorganic nutrients (Trench 1979).
Dependence on zooxanthellae led coral reefs to thrive in shallow waters of the
tropics, which are characterized by high levels of solar radiation. Reef-building corals
harvest solar radiation efficiently while meeting the physiological challenge of safely
disposing excess excitation energy (Hoegh-Guldberg and Jones 1999; Brown et al. 1999).
Ultraviolet (UV) radiation (280-400 nm) has been reported to have a variety of damaging
effects on marine organisms (Jokiel 1980), including corals and their dinoflagellate
symbionts (Shick et al. 1996). Reduced growth rates, chlorophyll a, carbon:nitrogen
ratios, photosynthetic oxygen evolution and ribulose bisphosphate carboxylase/oxygenase
(Rubisco) activity have been documented in cultured Symbiodinium spp. (Lesser 1996).
The host and at least some of its symbionts produce UV absorbing compounds, known as
mycosporine-like amino acids (MAAs), and possess several antioxidant systems to cope
with UV radiation (Shick et al. 1996 – for review).
Photosynthetically active radiation (PAR) can be potentially dangerous to
photosynthetic organisms. Above saturating levels, light energy is potentially detrimental
to the photosynthetic apparatus (Long et al. 1994). Reduction in the efficiency in
processing absorbed light has been termed photoinhibition (Kok 1956). Photoinhibition
can be described as dynamic and chronic depending on the rate of recovery of the
photosynthetic efficiencies. Dynamic photoinhibition (photoprotection) represents a
rapidly reversible downregulation of photosystem II (PSII) quantum yield and is
correlated with the thylakoid lumenal acidification occurring during light-dependent
electron transport. Chronic photoinhibition (photodamage) is primarily associated with
damage to PSII, particularly the D1 protein of this reaction center.
Photoprotective mechanisms in reef-building corals span a wide range of
phenomena from both partners. At the host level, these include behavioural responses
such as tissue retraction (Brown et al. 1994, 2002), fluorescent proteins (Salih et al.
2000), tissue thickness and colony morphology (Loya et al. 2001). The coral symbiotic
algae dissipate heat by the reversible inter-conversion of the xanthophylls diadinoxanthin
and diatoxanthin (Brown et al. 1999). To deal with downstream processes both the coral
host and the algae possess a range of antioxidant enzymes (Lesser 1997).
Originally Symbiodinium (Freudenthal 1962) was believed to be a monotypic
genus. Over the last two decades, Symbiodinium has been revealed to be highly diverse at
the biochemical, physiological and morphological levels (Blank and Trench 1985; Trench
and Blank 1987; Lajeunesse and Trench 2000). Recently molecular genetic studies using
restriction fragment length polymorphism (RFLP) analysis of the small subunit ribosomal
DNA (rDNA) have identified several clades A, B, C, D, E, F and G (Rowan 1998; Toller
et al. 2001; Loh et al. 2002) of zooxanthellae. Analysis based on internal transcribed
spacer 2 (ITS2) region using denaturing gradient gel electrophoresis (DGGE) has futher
subdivided the clades (Lajeunesse 2001, 2002). Clade A zooxanthellae have been
reported to dominate in shallow water corals while C and B prevail in deeper corals
(Rowan et al. 1997). Indeed, only clade A zooxanthellae in culture were found to produce
MAAs (Banaszak et al. 2000) and thus seems suitable for high light environments.
This paper investigates coral surface skeletal morphology and zooxanthellae types
in the plate growth form of the coral Montipora capitata inhabiting low and high-light
environments in the field. By using the technique of pulse-amplitude-modulation (PAM)
fluorometry the photosynthetic performances of the in hospite (within host tissue)
zooxanthellae from the two light environments were also examined.
Methods
Study site and sample collection
Two sites, the Lighthouse and Boatcove on Coconut Island, Hawaii, were chosen, and
low and high-light stations within each site were established using the underwater light
meter (LI1400) placed next to the coral colonies sampled. Low and high-light
environments represented shaded (sheltered) and exposed (open) areas inhabiting the
plate form of Montipora capitata. Temperature was also recorded over 1-3 days at the
stations using the I-button Thermochron (Model, 0036 A3 DS1921L-F52). Coral
fragments from four colonies from each site for both low and high light environments
were collected and maintained in running seawater table until examined.
Water motion was also determined at both sites for each station using the method
of Jokiel and Morrissey (1993). Slow-dissolving (S-type) clods, prepared from a mixture
of powdered plastic resin glue and wall patching compound and mounted on plastic
cards, were supplied by Dr. Jokiel. Briefly, fifteen clod cards were soaked in seawater for
24 h and weighed after rinsing with fresh water and blotting. The initial weight was
recorded. Three clod cards were kept in a bucket of still seawater in shade as control and
the rest were set at the sites with three replicates at each station. The clod cards were
reweighed after 24 h. The diffusion factor, which represents the ratio of weight loss in the
experimental blocks to weight loss in the calm water (control), was determined.
Surface skeletal morphology examination
The collected coral fragments from the low and high-light regimes from both sites were
observed under a binocular microscope. The number of tuberculae over a given surface
area (normalized to cm2) was quantified and digital pictures were also taken through the
binocular microscope. Tuberculae are defined as the projections of coenosteum on the
surface of many species of Montipora that are more than a corallite in width (Veron
2000).
Zooxanthellae isolation and DNA extraction
Coral pieces (2 x 2 cm2) from the respective collected colonies were airbrushed with
filtered seawater (FSW) (0.45 m) in small zip-lock bags. The blastate / slurry was
transferred to 15 ml tubes and centrifuged at 800 g for 5 min and washed with FSW
twice. The algal pellet was transferred to 1.5 ml microfuge tubes, volume made up to 1.5
ml with DMSO, and incubated overnight at 25oC.
The algae, preserved in the DMSO buffer, were centrifuged at 14,000 rpm for 5
min. To wash the excess DMSO 500 l TE was used. Centrifugation and washing with
TE was done, and the remaining fluid was dried using a vacuum suction system. Agal
DNA was extracted using Promega’s Wizard Genomic DNA Extraction Kit. 20-40 mg
(20-40 l) of the pellet was placed in a new 1.5 ml microfuge tube containing 250 l
sterile glass beads (0.5 mm) and 600 l nuclei lysis buffer. These tubes were then placed
in a bead-beater and beating was done for 3 min. They were then placed in water bath at
65oC for 5-10 min.
Three l of Proteinase K (20 mg/ml) was added and vortexed for 2-3 s. Tubes
were incubated at 65oC for at least 1 h with 2-3 s vortexing every 15-20 min. 1 l RNAse
(4 mg/l) was added and the extracts were incubated at 37oC for 15-20 min. 250 l protein
precipitation solution was added to each sample and vortexed for 5 s. The samples were
placed on ice for 15 min. The samples were centrifuged at 14,000 rpm for 10 min.
Six hundred l of clear supernatant was placed in a new 1.5 ml microfuge tube
containing 700 l isopropanol (100%) and 50 l 3 M sodium acetate. After gently mixing
the contents, samples were placed on ice for 10 min and then centrifuged at 14,000 rpm
for 10 min. The isopropanol was aspirated, and 500 l 70% ethanol was added. The
tubes were gently shaken, followed by a 5 min centrifugation at 14,000 rpm. The ethanol
was carefully aspirated and the sample was dried in the fume hood overnight.
ITS amplification and denaturing gradient gel electrophoresis (DGGE) analysis
The ITS2 region (330-360 bp) primers (modified from Lajeunesse and Trench 2000)
were used for PCR amplification. An internal primer “ITSintfor2” (5 GAATTGCAGA
ACTCCGTG-3) annealing to a conserved region of the 5.8S rDNA and pairing with the
conserved 3 flanking primer, ITS reverse, was used. The end primer, called the
“ITS2CLAMP” with a 39 bp GC clamp (underlined) (5 CGCCCGCCGC
GCCCCGCGCC CGTCCCGCCG CCCCCGCCC GGGATCCATA TGCTTAAGTT
CAGCGGGT-3) was employed. Reactions were carried out on a BioRad Thermal Cycler
using a “touchdown” amplification protocol with annealing conditions (Lajeunesse 2002)
with annealing conditions 10oC above the final annealing temperature of 52oC. A 1oC in
annealing temperature was allowed for every two cycles. Following 20 cycles the 52oC
annealing temperature was maintained for 15-18 further cycles.
The amplicons from the different specimens were run on a gradient gel. All
products were loaded with a 2% Ficoll loading buffer (2% Ficoll-400, 1.0 mM tris-HCl
pH 7.8, 1 mM EDTA, 1% bromophenol) onto an 8% polyacrylamide denaturing gel
(gradient of 3.15 M urea / 18% deionized formamide to 5.6 M urea / 37% deionized
formamide). Separation by electrophoresis was carried out at 160 V for 9.5 h at a constant
temperature of 60oC. The gel was stained with Syber Green (Molecular Probes, Eugene,
OR.) for 25 min in darkness and visualized with UV transillumination (BioRad
SyncMaster 753DF).
Chlorophyll a fluorescence measurements
Chlorophyll a fluorescence parameters were measured using a pulse-amplitudemodulated (PAM) underwater fluorometer (DIVING-PAM: Walz, Germany) on in
hospite zooxanthellae, that is, zooxanthellae within the tissues of coral fragments held in
seawater in a 300 ml containers. Each sample was illuminated for 5 min at a saturating
light intensity of 125 mol quanta m-2 s-1 using the internal actinic light source followed
by a 30 s dark period, prior to an automatic series of measurements using the internal
actinic (photosynthesis-inducing) light source. The initial fluorescence, Ft, was measured
by applying a weak pulsed light (< 1 mol quanta m-2 s-1 PAR) and a saturating pulse
(8000 mol quanta m-2 s-1) was used to determine the maximum fluorescence, Fm. The
change in fluorescence (Fv) caused by the saturating pulse, in relation to the maximal
fluorescence yield (Fm), has been shown to be a good measure of effective quantum
yield, Y (Genty et al. 1989).
The ratio, Y, is a measure of the effective quantum yield of PSII in an illuminated
sample. Since electrons leading to CO2 reduction in the dark reactions of photosynthesis
are derived from the splitting of water in PSII, photosynthetic electron transport rate
(ETR) may be estimated from the effective quantum yield. Thus, ETR is expressed by the
effective quantum yield (Y) x PFD x FA x 0.5, where PFD is the photosynthetic flux
density of photosynthetically active radiation (PAR, 400-700 nm), FA is the fraction of
the incident light absorbed by the living photosynthetic tissue, and an assumption is made
that two photons are necessary to produce one high energy electron.
In the present study, the actual actinic light levels that reached the measured coral
surface were determined using a LICOR lightmeter (Model, LI1400). The optic fibre
head of the fluorometer was fixed at an adjusted distance of 13 mm from the surface of
the specimens and the water depths was maintained constant for all the measurements.
An FA, used instead of the standard “ETR-factor” for estimating absolute values of ETR
(mol electrons m-2 s-1), of 0.036 (used by Beer et al. 1998 for the coral Favia favus) was
employed to calculate the ETRs. The rapid light curves (RLCs) for the in hospite
zooxanthellae were fitted by third order polynomials as this order gave the best r2 values.
The gain setting on the fluorometer was set to two for all measurements.
Results
Environmental parameters
The physical parameters monitored in this study were light intensity and temperature (2-3
days period at each station) and water motion (estimated as diffusion factor over 24 h).
Fig. 1 shows the daily variations in light intensity in the low and high-light characterized
stations. At both sites (Fig. 1A, B) the corals inhabiting the high-light environment
experienced at least 28 fold higher maximum light levels than those living in the lowlight one. The temperature in the shaded stations at both the Lighthouse and Boatcove
ranged between 25.0 - 25.5oC. The exposed stations had temperature ranges of 24.5 -
25.5oC and 23.0 - 25.5oC at the Lighthouse and Boatcove, respectively. Diffusion factor
did not differ between stations (Lighthouse - 7.17  0.84, 9.18  0.33; Boatcove – 4.78 
0.18, 5.03  0.56, mean  SD for low and high-light environments, respectively) but were
twice higher at the Lighthouse than at the Boatcove.
Coral surface morphology in different light environments
The coral fragments collected from the two light environments at two different sites had
different surface morphologies (Fig. 2). Fig. 3. shows the presence or absence of
tuberculae in the low and high-light environment coral samples. The high-light corals had
significantly higher number of tuberculae per cm2 (Table 1) than those surviving in the
low light regime.
DGGE profile of zooxanthellae in corals from low and high light environments
Fig. 4 depicts the ITS2 typing of the zooxanthellae from Montipora capitata living under
different light regimes. A distinctive band pattern sometimes accompanied by faint
“background” bands typically characterized the DGGE profiles from the isolated
zooxanthellae from the coral under investigation. The profiling reveals that the type of
zooxanthellae is the same in shade or light collected M. capitata and is representative of
the C17 ITS2 type (Lajeunesse pers. com.).
Photosynthetic performance of in hospite zooxanthellae from the different light
environments studied
The RLCs obtained by employing PAM fluorometer is representative of P-I
(photosynthesis versus irradiance) curves. The RLCs are shown in Fig. 5 for in hospite
zooxanthellae surviving in low and high-light regimes. The maximum electron transport
rate (ETRmax), light level at which photosynthesis saturates (Ek) and light utilization
efficiency () did not differ significantly when corals from the two light regimes were
compared. However, an earlier and more pronounced photoinhibition was observed in the
low-light corals when compared to the high-light ones.
Discussion
The physical environment influences both the coral animal host, at least at the skeletal
deposition level (Buddemeier and Kinzie 1976) and their endosymbiont types (Rowan
and Powers 1992; Baker 2002; Lajeunesse 2002). In this study, temperature, depth and
water motion were not significantly different between the low and high-light chosen
environments within the given site. Therefore, comparisons between stations within each
site can mostly be attributed to the effect of light environments on the coral Montipora
capitata.
Previous works suggest that physical environment can affect the coral skeletal
formation and growth and soft tissue. Both light (Houck et al. 1977) and water motion
(Jokiel 1978) can influence growth rates and form. Changes in shape of colonies from
hemispherical to plate-like morphologies with increasing depth have been reported in
Montastrea annularis (Goreau 1963; Dustan 1975), Porites asteroides (Roos 1967) and
Synaraea convexa (Jaubert 1977). These authors considered light to be the controlling
agent. Polyp density has been reported to decrease with depth (Lasker 1981; Davies
1980). In this study polyp density appeared to be lower (observation, not quantified) in
corals from low-light environment. Interestingly, tuberculae density was found to be high
in high-light environment specimens. It should be noted that almost no tubercula was
formed on the lower surface of either the low or high-light environment coral specimens.
One might be tempted to interpret this as tuberculae formation being a light-induced
phenomenon. Olivier et al. (1983) reported more rapid branch initiation in Acropora
formosa from shallow sites.
To verify the possibility that the tuberculae are assuming a photoprotective role,
the photosynthetic responses of the corals from low and high-light regimes were also
measured. It is well established that light and shaded adapted corals (Jones and HoeghGuldberg 2001), cultured Symbiodinium spp. (Iglesias-Prieto and Trench 1994) and other
marine diatoms (Mock and Kroon 2002) have different characteristic photosynthetic
responses. However, there were no significant differences in the conventional
photosynthetic parameters such as the maximum electron transport rate (ETRmax), light
level at which photosynthesis saturates (Ek) and light limited slope of the rapid light
curves representing the light utilization efficiency () between the low and high-light
collected coral specimens. This is indicative that the low light levels in the shaded
environment were still sufficient to maintain the photosynthesis level required for the
symbiosis to exist and the coral to survive. On the other hand, the high-light environment
was not detrimental to the zooxanthellae as the light level was probably buffered to
protect the algae from chronic damage. Another photosynthetic feature revealed here is
the earlier and more pronounced photoinhibition observed in the low-light environment
collected corals with very low density of tuberculae. This most probably suggests a role
of tuberculae in photoprotection.
Denaturing gradient gel electrophoresis analysis revealed no difference in the
ITS2 types of zooxanthellae harbored by the corals from the two light regimes under
investigation. This is in contrast with the depth zonation of zooxanthellae reported in
some studies (Rowan et al. 1997; Baker 2001). However, this result is not surprising, as
the zooxanthellae in Montipora capitata are known to be vertically transmitted and might
exhibit a close system as far as change of endosymbionts is concerned. Furthermore, the
ETRmax, Ek and  did not vary significantly between the low and high-light corals
suggesting that the zooxanthellae are receiving enough light for maintaining their
photosynthetic performances.
In summary, this study highlights the role of the host in terms of increasing
number of tuberculae on the coral surface in order to photoprotect the zooxanthellae from
photoinhibition. The algae ITS2 typing and their acclimatory photosynthetic
characteristics such as ETRmax, Ek and  showed no difference between the low and highlight environment corals. Further reciprocal transplantation experiments are warranted if
the role of light in inducing the formation of tuberculae is to be understood thoroughly.
Acknowledgments
The author is thankful to the staffs of the Hawaii Institute of Marine Biology (HIMB),
University of Hawaii and Edwin Pauley Foundation for the opportunity granted to attend
the 2002 summer course. RB is greatly indebted to the usual help and support of his
supervisor, Dr M Hidaka. Special thanks are due to Dr Todd Lajeunesse for his generous
expertise, experience and technical help in DGGE analysis. Appreciation is also extended
to Drs PL Jokiel, J Stimson and MP Lesser for providing clod cards, I-buttons and Diving
PAM, respectively. Much gratitude is also due to the student participants of the summer
program. This manuscript also benefited from comments from Drs IM Yakovleva and EA
Titlyanov. The invaluable advice and excellent laboratory facilities provided by Drs
Fenny Cox, Robert A Kinzie III, Teresa Lewis and Deb Shearman throughout the course
are greatly appreciated. This work was partly funded by grant-in-aid for Joint Research
Projects of Japan-US Bilateral programs from Japanese Society for the Promotion of
Science (Principal investigator: Dr K Sakai) and a scholarship from the Ministry of
Education, Culture, Sports, Science and Technology, Japan. Corals were collected under
the scientific collection permit #2003-06 to HIMB.
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Table 1. Counts of tuberculae normalized to cm2 area in the plate form of Montipora
capitata inhabiting low and high light environments. Data represent mean  SD (n = 4
colonies). Asterisk indicates significant difference (Student t-test, P <0.001) when the
high-light data were compared with the low-light ones at each site.
Low-light
High-light
Lighthouse
0.14  0.16
14.20  2.19*
Boatcove
0.07  0.11
9.83  1.59*
Figure captions
Fig. 1. Diel changes in underwater light intensity at Lighthouse (A) and Boatcove (B) for
exposed and shaded stations. The underwater LI-COR (LI 1400) probe was placed next to
the colonies under investigation. Monitoring was done for three and two days at
Lighthouse and Boatcove, respectively.
Fig. 2. Surface morphology of the plate form of Montipora capitata from Boatcove and
Lighthouse. The upper coral fragments were collected from the high-light environment
while the lower ones from the low-light environment.
Fig. 3. Surface morphology observations under the binocular microscope using low-light
(A) and high-light (B) specimens. C and D show absence and presence of tuberculae in
corals collected from low-light and high-light environments, respectively.
Fig. 4. Denaturing gradient gel electrophoresis (DGGE) profile based on the amplified
ITS2 region of the ribosomal DNA of zooxanthellae isolated from Montipora capitata
inhabiting either low or high light environments. LHS = Lighthouse shaded; LHE =
Lighthouse exposed; BHS = Boatcove shaded; BHE = Boatcove exposed; C17 = ITS2
type of zooxanthellae belonging to clade C (identified by Dr. Lajeunesse).
Fig. 5. Rapid light curves for the coral Montipora capitata collected from low (dotted
line) and high light (solid line) environments. Curves were fitted by third order
polynomials with r2 > 0.96. Data points represent mean  SD (n = 4 colonies).
FIGURE 1
-2 -1
Light intensity ( mol quanta m s )
A
1600
1400
1200
1000
800
600
400
200
0
12:00
0:00
12:00
0:00
12:00
0:00
Time of day (h)
12:00
0:00
12:00
FIGURE 2
Boatcove
High-light
Low-light
Lighthouse
FIGURE 3
A
C
B
D
FIGURE 4
LHS
LHE
BHS
BHE
C17
FIGURE 5
Electron transport rate
1
0.8
0.6
0.4
0.2
0
0
100
200
300
400
500
600
Light intensity (mol quanta m-2 s -1)
700