The effects of environmental history and thermal stress on coral physiology and immunity

Abstract

Rising ocean temperatures can induce the breakdown of the symbiosis between reef building corals and Symbiodinium in the phenomenon known as coral bleaching. Environmental history may, however, influence the response of corals to stress and affect bleaching outcomes. A suite of physiological and immunological traits was evaluated to test the effect of environmental history (low vs. high variable pCO2) on the response of the reef coral Montipora capitata to elevated temperature (24.5 °C vs. thermal ramping to 30.5 °C). Heating reduced maximum photochemical efficiency (Fv/Fm) and chlorophyll a but increased tissue melanin in corals relative to the ambient treatment, indicating a role of the melanin synthesis pathway in the early stages of thermal stress. However, interactions of environmental history and temperature treatment were not observed. Rather, parallel reaction norms were the primary response pattern documented across the two temperature treatments with respect to reef environmental history. Corals with a history of greater pCO2 variability had higher constitutive antioxidative and immune activity (i.e., catalase, superoxide dismutase, prophenoloxidase) and Fv/Fm, but lower melanin and chlorophyll a, relative to corals with a history of lower pCO2 variability. This suggests that reef environments with high magnitude pCO2 variability promote greater antioxidant and immune activity in resident corals. These results demonstrate coral physiology and immunity reflect environmental attributes that vary over short distances, and that these differences may buffer the magnitude of thermal stress effects on coral phenotypes.

Full text

Vol.:(0123456789)
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Marine Biology (2018) 165:56
https://doi.org/10.1007/s00227-018-3317-z
ORIGINAL PAPER
The eects ofenvironmental history andthermal stress oncoral
physiology andimmunity
ChristopherB.Wall1 · ContessaA.Ricci2· GraceE.Foulds1· LauraD.Mydlarz2· RuthD.Gates1· HollieM.Putnam3
Received: 16 November 2017 / Accepted: 16 February 2018
© Springer-Verlag GmbH Germany, part of Springer Nature 2018
Abstract
Rising ocean temperatures can induce the breakdown of the symbiosis between reef building corals and Symbiodinium in
the phenomenon known as coral bleaching. Environmental history may, however, influence the response of corals to stress
and affect bleaching outcomes. A suite of physiological and immunological traits was evaluated to test the effect of environ-
mental history (low vs. high variable pCO2) on the response of the reef coral Montipora capitata to elevated temperature
(24.5°C vs. thermal ramping to 30.5°C). Heating reduced maximum photochemical efficiency (Fv/Fm) and chlorophyll a
but increased tissue melanin in corals relative to the ambient treatment, indicating a role of the melanin synthesis pathway
in the early stages of thermal stress. However, interactions of environmental history and temperature treatment were not
observed. Rather, parallel reaction norms were the primary response pattern documented across the two temperature treat-
ments with respect to reef environmental history. Corals with a history of greater pCO2 variability had higher constitutive
antioxidative and immune activity (i.e., catalase, superoxide dismutase, prophenoloxidase) and Fv/Fm, but lower melanin
and chlorophyll a, relative to corals with a history of lower pCO2 variability. This suggests that reef environments with high
magnitude pCO2 variability promote greater antioxidant and immune activity in resident corals. These results demonstrate
coral physiology and immunity reflect environmental attributes that vary over short distances, and that these differences may
buffer the magnitude of thermal stress effects on coral phenotypes.
Introduction
The mutualistic symbiosis between scleractinian corals and
dinoflagellates of the genus Symbiodinium underpins the
function of hermatypic corals and their capacity to engineer
tropical reef ecosystems (Putnam etal. 2017). Environmen-
tal disturbances destabilize this symbiosis and reduce the
abundance of Symbiodinium cells and/or their photosyn-
thetic pigments within coral tissues; a stress response called
coral bleaching (Coles and Jokiel 1978). Elevated seawater
temperatures have driven three global coral bleaching events
to date (Hoegh-Guldberg etal. 2017), and ocean warming
and the frequency of bleaching-level stress are predicted to
increase as climate change intensifies (Heron etal. 2016;
Hughes etal. 2017). While corals have persisted through
considerable environmental change in the geologic record
(Pandolfi and Kiessling 2014), the magnitude and rate of
change in the thermal and chemical properties of seawater
during the Anthropocene is unprecedented (Zeebe 2012;
IPCC 2014; Hubbard 2015).
The response of corals to thermal stress is influenced
by physical conditions that precede (Brown etal. 2002a;
Middlebrook etal. 2008; Carilli etal. 2012; Guest etal.
2012; Ainsworth etal. 2016) and/or co-occur with elevated
temperatures (Coles and Jokiel 1978; Dunne and Brown
2001; Nakamura and van Woesik 2001; Jokiel and Brown
2004; Anthony etal. 2008; Wiedenmann etal. 2012). It
is recognized that organisms are equipped with diverse
biochemical mechanisms to acclimate and adapt to physi-
ological stress (Hochachka and Somero 2002), and in cor-
als, evidence supports the role of the coral animal (Kenkel
Responsible Editor: R. Hill.
Reviewed by C. Palmer and an undisclosed expert.
* Christopher B. Wall
cbw0047@gmail.com
1 University ofHawai‘i atMānoa, Hawai‘i Institute ofMarine
Biology, Kaneohe, HI96744, USA
2 Department ofBiology, University ofTexas atArlington,
Arlington, TX76019, USA
3 Department ofBiological Sciences, University ofRhode
Island, Kingston, RI, USA

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etal. 2013a) and Symbiodinium (Levin etal. 2016) in con-
fronting environmental challenges (Edmunds and Gates
2008; Hume etal. 2016; Palumbi etal. 2014). For instance,
corals experiencing thermal (Lesser 2004; Fitt etal. 2009;
Kenkel etal. 2011, 2013b) and/or photo-stress (Brown
etal. 2002b) can mitigate cellular damage by up-regulating
stress proteins (i.e., fluorescent and heat-shock proteins)
(Lesser 2004; Palmer etal. 2009; Louis etal. 2017). Coral
immunity and pathogen defense mechanisms (e.g., mela-
nin synthesis pathway) (Söderhäll and Cerenius 1998) are
also dynamically regulated in response to bleaching stress
(Mydlarz etal. 2009). Indeed, maintaining high baseline
immunity and tissue-protective properties (e.g., antioxida-
tive enzymes) may represent a conserved mechanism of
coral physiological resilience to both disease and thermal
stress (Weis 2008; Palmer etal. 2010; Louis etal. 2017).
Rising concentrations of carbon dioxide (pCO2) and
other greenhouse gases are driving global climate change
by increasing air and ocean temperatures (IPCC 2014).
In addition, the dissolution of atmospheric CO2 in the
upper ocean is disrupting seawater carbonate chemistry
and causing ocean acidification (OA), which threatens the
net calcification of coral reef ecosystems (Andersson and
Gledhill 2013). Exposure to elevated pCO2 also has the
potential to influence coral immunity and the response
of corals to warming temperatures (Anthony etal. 2008;
Kaniewska etal. 2012). For instance, pCO2 can exacerbate
thermal stress effects and cause bleaching in some corals
(Anthony etal. 2008; but see Wall etal. 2014; Noonan
and Fabricius 2016). Additionally, corals in experimen-
tally elevated pCO2 conditions or from naturally high
pCO2-seeps show an upregulation of genes involved in
oxidative stress and innate immune pathways (Kaniewska
etal. 2012; Kenkel etal. 2017).
Natural field settings where elevated pCO2 conditions
are persistent (Fabricius etal. 2011; Albright etal. 2015;
Padilla-Gamiño etal. 2016; Kenkel etal. 2017), or dynamic
in nature (Drupp etal. 2013), can provide insight into the
consequences of high pCO2/low pH on marine taxa not
apparent in short-term laboratory experiments (Calosi etal.
2013; Noonan and Fabricius 2016). Leveraging natural
field settings with unique seawater properties can, there-
fore, clarify the influence of pCO2 history on coral physi-
ology and thermal stress responses (Noonan and Fabricius
2016). Within Kāne‘ohe Bay (windward O‘ahu, Hawai‘i)
a combination of factors (e.g., physical forcing, seawater
residence time, watershed and oceanic biogeochemistry)
(Lowe etal. 2009; Drupp etal. 2011, 2013; Shamberger
etal. 2011) has created regions where corals are exposed to
seawater pCO2 projected to occur under end-of-the-century
climate change scenarios (van Vuuren etal. 2011). As such,
Kāne‘ohe Bay provides an ideal natural setting to address the
hypothesis that environmental history—specifically, regimes
of contrasting pCO2 variability (Drupp etal. 2011, 2013)—
alters the biology of reef corals and their response to stress.
The dynamic interplay between multiple stressors, envi-
ronmental history and physiological acclimatization shapes
reef resilience in varying ways. The goal of this study was to
test the interaction of environmental history in the context of
pCO2 variability and short-term thermal stress on the physi-
ological, photochemical, and immunological responses of
corals from two Kāne‘ohe Bay reefs with contrasting pCO2
conditions. Considering the potential for elevated pCO2 to
negatively influence coral performance and cause bleach-
ing, we tested the hypothesis that corals from environments
with a history of high variable pCO2 would show greater
sensitivity to thermal stress by exhibiting greater declines in
photochemical efficiency, photopigment concentrations and
Symbiodinium densities relative to corals from environments
with a history of low variable pCO2. We also expected corals
from high variable pCO2 environments to display increased
antioxidative activity as a mechanism to mitigate cellular
damage (Weis 2008), as well as greater immune activity
measured by elevated melanin synthesis pathway activity
(prophenoloxidase and melanin).
Materials andmethods
Study site description
Reef selection was driven by previous characterization
of the physical and chemical conditions occurring within
the reef–lagoon system of Kāne‘ohe Bay, O‘ahu, Hawai‘i
(21°26′06.0″N, 157°47′27.9″W) (Lowe etal. 2009; Drupp
etal. 2011, 2013; Shamberger etal. 2011). The hydrody-
namics of Kāne‘ohe Bay are highly heterogeneous due to
different physical forcing (i.e., wave, wind, tidal) among bay
regions (Lowe etal. 2009). In the southern lagoon (zone 6,
sensu Lowe etal. 2009), geographic isolation and resist-
ance to wave-driven forcing reduce seawater mixing and pro-
duce prolonged seawater residence times (ca. 30–60days).
Conversely, in the central lagoon (zone 5, sensu Lowe etal.
2009) seawater residence times are reduced (ca. 10days)
due to greater wave-driven forcing and oceanic influences
(Lowe etal. 2009). The nexus of these physical factors and
biological processes (i.e., photosynthesis/respiration, calcifi-
cation/dissolution) produce distinct pCO2 conditions within
Kāne‘ohe Bay (Drupp etal. 2011, 2013; Shamberger etal.
2011). The NOAA Pacific Marine Environmental Labo-
ratory (PMEL) and the University of Hawai‘i Coral Reef
Instrumented Monitoring and CO2-Platform (CRIMP) buoys
provide high-resolution time-series data for water column
carbonate chemistry and pCO2, which have been applied to
evaluate the relationship between physical forcing, nutrient
input, and reef metabolism on air–sea CO2 exchanges on

Marine Biology (2018) 165:56
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reefs across O‘ahu, including Kāne‘ohe Bay (Fig.1) (see
De Carlo etal. 2007; Drupp etal. 2011, 2013; Shamberger
etal. 2011). These data reveal water column pCO2 in south-
ern Kāne‘ohe Bay (buoy: CRIMP-1, Sabine etal. 2012, or
CRIMP–CO2 buoy sensu Drupp etal. 2011) and adjacent to
the barrier reef (buoy: CRIMP-2 buoy, sensu Drupp etal.
2013) are comparable (ca. 450μatm pCO2); however, the
range in pCO2 varies in these two regions of Kāne‘ohe Bay,
being 225–671μatm pCO2 at CRIMP-1 (i.e., low variable
pCO2) (Fig.2a) and 196–976μatm pCO2 at CRIMP-2 (i.e.,
high variable pCO2) (Fig.2b) (Drupp etal. 2011, 2013).
Reefs adjacent to CRIMP-1 and CRIMP-2 buoy deploy-
ments were identified as the sites for this study. Corals were
collected from an inshore fringing reef (21°25′36.8″N,
157°47′24.0″W) (Lilipuna) located in the southwestern
basin of Kāne‘ohe Bay, 350m south of Moku o Lo‘e [Coco-
nut Island and the Hawaiian Institute of Marine Biology
(HIMB)] and proximate to CRIMP-1 (Drupp etal. 2011),
and an inshore patch reef (21°27′08.6″N, 157°48′04.7″W)
(Reef 14) in the central lagoon of Kāne‘ohe Bay and adja-
cent to CRIMP-2 (Drupp etal. 2013) (Fig.1). Hereafter,
the two reefs where corals were collected will be referred to
as ‘low variable pCO2 Lilipuna’ (LV–Lilipuna)—adjacent
to CRIMP-1, experiencing prolonged seawater residence
−157.82
157.82°
W1
57.78°
21.41 21.41°
21.44°
21.47° N
LV−Lilipuna
HV−Reef 14
CRIMP−1
CRIMP−2
Moku o Lo'e
Fig. 1 Map of Kāne‘ohe Bay on the windward side of the island of
O‘ahu, Hawai‘i, USA, detailing locations of two reefs characterized
by an environmental history of low variable pCO2 (LV–Lilipuna) and
high variable pCO2 (HV–Reef 14), NOAA PMEL buoys (CRIMP-1
and CRIMP-2), and the Hawai‘i Institute of Marine Biology (Moku
o Lo‘e)
200 400 600 800 1000
Air
Seawater
Sep '06 Nov '06 Jan '07 Mar '07 May '07 Jul '07Sep '07 Nov '07 Jan '08 Mar '08 May '08
200400 600800 1000
pCO2(µatm)
Jun '08 Aug '08 Oct '08 Dec '08 Feb '09 Apr '09 Jun '09 Aug '09 Oct '09 Dec '09
Date
pCO2(µatm)
a
CRIMP-1
b CRIMP-2
Fig. 2 Concentrations of carbon dioxide (μatm pCO2) in seawater and
air at CRIMP-1 (upper panel) and CRIMP-2 (lower panel) moored at
two locations within Kāne‘ohe Bay, O‘ahu, Hawai‘i, USA (see Fig.1)
(Data: NOAA PMEL; Drupp et al. 2011, 2013; Sabine et al. 2012;
Sutton etal. 2016)

Marine Biology (2018) 165:56
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56 Page 4 of 15
and low pCO2 flux—and ‘high variable pCO2 Reef 14’
(HV–Reef 14)—adjacent to CRIMP-2, experiencing short
seawater residence and high pCO2 flux.
Environmental monitoring
Seawater pCO2 conditions proximate to each reef (LV–Lili-
puna, HV–Reef 14) were sourced from quality-controlled,
publically available NOAA PMEL-CRIMP CO2-Platform
moored buoys (https ://www.pmel.noaa.gov/co2/story /
Coral +Reef+Moori ngs) (Sabine etal. 2012; Sutton etal.
2016), using two deployment periods: CRIMP-1 (near
LV–Lilipuna) 25 Nov 2005–16 Jun 2007, and CRIMP-2
(near HV–Reef 14) 11 Jun 2008–12 Jan 2010. During the
present study’s experimental period (Jan–Apr 2014), Hobo
loggers (Onset Computer Corp., Bourne, Massachusetts)
cross-calibrated to a certified digital thermometer (5-077-
8,±0.05°C, Control Company, Webster, Texas) recorded
temperatures at each reef site at the depth of coral collec-
tion (<1m). Separately, a comparison of light availabil-
ity at LV–Lilipuna and HV–Reef 14 was performed (Oct
2014–Dec 2014) at<1m for each reef using Odyssey pho-
tosynthetic irradiance loggers (Dataflow Systems Limited,
Christchurch, New Zealand) cross-calibrated to a Licor
(LI-1400, Lincoln, Nebraska) equipped with a cosine quan-
tum sensor (LI-192) (Long etal. 2012). While collections
of environmental data (i.e., pCO2, temperature, PAR) are
temporally distinct and reefs may not experience seawater
with identical carbonate chemistry as measured at buoys,
collectively, these data are useful in describing trends in
environmental characteristics among the two reef locations
and their relationship to coral performance.
Coral collection andlaboratory treatments
Fifty M. capitata (Dana 1846) branch tips (ca. 4cm in
length) were collected from each reef on 5 February
2014 (State of Hawai‘i Department of Land and Natu-
ral Resources, Special Activity Permit 2013-47); accord-
ingly, holobiont biomass should be considered seasonally
acclimated to Kāne‘ohe Bay winter conditions (Fitt etal.
2000). Fragments were transported in seawater to HIMB
and epoxied to plastic bases using Z-spar A788 splash zone
compound in a flow-through water table. One day after
collection, corals were transferred into two custom-built
experimental flow-through aquaria (50L; Aqualogic, Inc.,
North Haven, Connecticut) receiving sand-filtered seawater
from Kāne‘ohe Bay at a rate of ca. 0.2Lmin−1 and main-
tained at ambient conditions of 36 salinity and ca. 24.5°C.
After 1week of acclimation to laboratory conditions, corals
(N=100) were randomly allocated to four flow-through
aquaria (50L) (two replicate tanks treatment−1) at a density
of 25 fragments tank−1.
Seawater temperatures in each tank were independently
regulated using a combination of 100W submersible heat-
ers and a programmable solenoid controller that inde-
pendently regulated the delivery of chilled water through
an in-line mixing column (Multi Temp MT-1 Model
#2TTB3024A1000AA, Aqualogic) receiving tank seawater.
Temperature treatments represented ambient temperature
conditions (24.5°C: Ambient) for January–February 2014
(NOAA 2017) and a heated treatment gradually ramped to
elevated temperatures (30.5°C: Heated) (Fig.3). Tempera-
ture ramping lasted 7days and increased at a rate of ca.
0.75°Cday−1. Corals were maintained at 30.5°C for 2days,
which is near the upper thermal limit of Hawaiian reef cor-
als (Coles etal. 1976; Coles and Jokiel 1978). The ramp-
ing regime was comparable to other studies (Middlebrook
etal. 2010) and was implemented to avoid acute heat shock,
ensuring observation of progressive heating effects on coral
performance. Corals were exposed to treatments from 11
to 19 February 2014. ANOVA confirmed the establishment
of two separate temperature treatments (F1,239=231.300,
P< 0.001); temperatures did not differ among replicate
heated tanks (F1,130=0.018, P=0.893) but replicate ambi-
ent tanks differed by 0.24°C (F1,107=71.578, P<0.001).
During the experiment, temperature was monitored
throughout the day using a certified digital thermometer
(Fisher Scientific 15-077-8, ±0.05°C, Hampton, New
Hampshire); photosynthetically active radiation (PAR) and
salinity were measured daily at three time points (10:00,
12:00, 16:00h) using a 4π-spherical quantum sensor (Li-
Cor) and a conductivity meter (Model 63, YSI Inc., Yellow
25
27
29
31
2468
Day of Treatment
Temperature (°C)
Ambient
Heated
Fig. 3 Raw temperature measurements from experimental treatments
representing an ambient (Ambient, gray symbols) and a progressively
warming condition (Heated, black symbols). Shaded regions for each
treatment indicate 95% confidence intervals of locally weighted least
squares regression

Marine Biology (2018) 165:56
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Page 5 of 15 56
Springs, Ohio), respectively. Light was supplied to each
tank by LED-lamps (Sol Super Blue, Aqua Illumination,
Ames, Iowa) on a 12-h light:12-h dark cycle and pro-
grammed to mimic diel changes in light intensities from
sunrise (06:00h) to sunset (18:00h) (Gibbin etal. 2015).
Photosynthetically active radiation (PAR) at the daily max-
imum (12:00h) was ca. 750μmol photons m−2s−1, and
each treatment tank received a mean (±SE, n=19) PAR of
452–467±54–57μmol photonsm−2s−1. PAR did not differ
among the four treatment tanks (F3,72=0.014, P=0.998)
or among days of the experiment (F6,69=0.894, P=0.505).
Physiological metrics
Pulse amplitude modulation (PAM) fluorometry was used
to measure temperature effects on the photochemical per-
formance of Symbiodinium spp. in hospite for all corals
on the 8th day of treatment exposure using a Diving-PAM
(Waltz, GmbH, Effeltrich, Germany). The Diving-PAM
was operated at a gain of 7, saturation intensity of 8, an
electronic signal damping of 2; under these conditions, the
signal to noise ratio was optimized and the minimum fluo-
rescence was stabilized at ca. 400–700 (arbitrary units). The
minimum (Fo) and maximum (Fm) fluorescence yield and
the maximum photochemical efficiency (Fv/Fm) of dark-
adapted PSII reaction centers were measured during the
day at 14:00h (which coincided with the period of peak
PAR exposure (Jones and Hoegh-Guldberg 2001) follow-
ing a 20-min dark acclimation period. Measurements were
obtained using the 5-mm diameter fiber-optic probe posi-
tioned 5mm above the surface of the coral tissue following
Fo stabilization. Following 9days of exposure, all corals
were immediately snap-frozen in liquid nitrogen and stored
at –80°C. A subset of corals were used for physiological
assays (Symbiodinium density, chlorophyll a extraction, and
total protein; n=10–13 treatment−1) and analyzed at HIMB.
Another subset of corals were used in immunological assays
(protein, melanin, prophenoloxidase, catalase, and superox-
ide dismutase; n=11–12 treatment−1) and analyzed at the
University of Texas at Arlington. Corals remained at –80°C
and were not thawed prior to analysis.
For physiological analyses, coral tissue was removed
from the skeleton using an airbrush and filtered seawa-
ter (0.7μm). The resulting coral tissue slurry was briefly
homogenized and aliquots taken for each response variable,
following Wall etal. (2017). The coral skeleton was placed
in 10% bleach solution and allowed to dry at 60°C before
measuring the surface area of the skeleton by the paraffin
wax-dipping technique (Stimson and Kinzie 1991). Sym-
biodinium densities were quantified by repeated cell counts
(n=6 sample−1) using a haemocytometer, and cell densi-
ties were standardized to coral surface area (cells cm−2).
Chlorophyll a was extracted by centrifuging the tissue slurry
(13,000rpm×3min) and isolating the alga pellet, followed
by adding 100% acetone and extracting at −20°C in dark-
ness for 36h. The pigment extract was measured spectro-
photometrically (λ=630 and 663nm) and chlorophyll a
concentrations were determined using equations for dino-
flagellates (Jeffrey and Humphrey 1975). Chlorophyll a was
standardized to surface area (μgcm−2) and to the density
of Symbiodinium cells (pg cell−1). Total protein (soluble
and insoluble) in the tissue slurry was measured using the
Pierce BCA (bicinchoninic acid) Protein Assay Kit (Pierce
Biotechnology, Waltham, Massachusetts). Solubilization of
protein was achieved by adding 1M NaOH to the tissue
slurry, heating at 90°C for 1h, and neutralizing to pH ca.
7.5 using 1N HCl. The total protein in three technical rep-
licates sample−1 was measured in a 96-well microtiter plate
(λ=562nm) against a bovine serum albumin standard curve
and standardized to coral surface area (mg protein cm−2).
Immunological assays
Coral immunology was assessed following previously
established protocols for protein extractions and enzymatic
assays (Mydlarz etal. 2009, 2010; Palmer etal. 2010, 2011a;
Mydlarz and Palmer 2011). Briefly, 3–4mL of coral tissue
slurry was obtained by airbrushing with coral extraction
buffer (100mM TRIS buffer+0.05mM dithiothreitol). The
resulting slurry was homogenized for 1min on ice using a
hand-held tissue homogenizer (Powergen 125, Fisher Sci-
entific, Waltham, Massachusetts). For melanin concentra-
tion estimates, 1mL of the tissue slurry was freeze-dried
for 24h using a VirTis BTK freeze-dryer (SP Scientific,
Warminster, Pennsylvania). The remaining slurry was cen-
trifuged at 4°C at 2500×g (Eppendorf 5810 R centrifuge,
Hamburg, Germany) for 5min to remove cellular debris,
and enzymatic assays were performed on aliquots of the
supernatant, representing a cell-free extract or soluble pro-
tein extract of the host coral. All assays were run in dupli-
cates on separate 96-well microtiter plates using a Synergy
HT multidetection microplate reader using Gen5 software
(Biotek Instruments, Winooski, Vermont). Protein concen-
trations were estimated using the RED660 protein assay (G
Biosciences, Saint Louis, Missouri) against a bovine serum
albumin standard curve.
Antioxidant prole
Antioxidant enzymes catalase (CAT) and superoxide dis-
mutase (SOD) were measured. CAT is monitored as a
change in absorbance after 25mM hydrogen peroxide is
added to crude protein extract and 50μL of 10mM PBS
(pH 6.0). CAT activity was estimated as the mM H2O2
scavenged min−1 mg protein−1. SOD activity was analyzed
using a commercially available kit (SOD determination

Marine Biology (2018) 165:56
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56 Page 6 of 15
kit #19160; Sigma-Aldrich, St. Louis, Missouri) following
manufacturer’s instructions and expressed as SOD activity
mg protein−1. SOD activity was estimated by comparing
the absorbance of samples at 450nm to a positive and
negative standard after incubating 10μL of crude protein
extract with manufacturer-provided reagents.
Melanin synthesis pathway
Prophenoloxidase (PPO) activity and melanin (MEL)
concentration per sample were used to study the mela-
nin synthesis pathways. PPO activity was determined by
incubating 20μL of protein extract and 50μL of 10mM
phosphate buffered saline (PBS) (pH 7.0) at room tempera-
ture with 20μL of trypsin (0.2mgmL−1 concentration)
for 30min. 20μL of 25mM l-DOPA (Sigma-Aldrich) was
then added as a substrate. PPO activity was estimated as
change in absorbance min−1mg protein−1. MEL concen-
tration was estimated using a weighed freeze-dried portion
of initial tissue slurry. Melanin was allowed to extract for
48h in 400μL of 10M NaOH after a brief period of bead-
beating with 1-mm glass beads. 65μL of extracted mela-
nin was used to determine endpoint absorbance at 495nm
and resulting values were standardized to a standard curve
of commercial melanin (Sigma-Aldrich) and calibrated to
μg melanin mg tissue−1.
Statistical analysis
Dependent variables were analyzed using a two-way linear
model using lme with temperature treatment (Ambient vs.
Heated) and reef environmental history (LV–Lilipuna vs.
HV–Reef 14) as fixed effects. Environmental data (light
and temperature) were analyzed using a linear mixed effect
model in the package lme4 (Douglas etal. 2015) with
site as a fixed factor and the repeated measure (sampling
time) as a random factor. Analysis of variance tables(lin-
ear models) and analysis of deviance tables (linear mixed
effect models) were calculated using Type-II sum of
squares with Satterthwaite approximation of degrees of
freedom using lmerTest (Kuznetsova etal. 2016). The
assumptions of analysis of variance were confirmed by
graphical inspection of residuals combined with Shap-
iro–Wilk’s test and Levene’s test and transformed where
assumptions of ANOVA were not met. Transformations
were selected using a Box–Cox power transformation
using the package MASS (Box and Cox 1964; Venables
and Ripley 2002). All analyses were performed in R, ver-
sion 3.3.0 (R Development Core Team 2016). Experimen-
tal data and R code to reproduce figures and analyses are
accessible on Zenodo (https ://zenod o.org/recor d/11750
34).
Results
Environmental data
During the experimental period (Jan–Apr 2014), temperature
did not differ among the two study sites (F1,12417=2.961,
P=0.085). Similarly, the daily mean temperature (ca.
24.6°C) (F1,65=0.137, P=0.713) and maximum daily tem-
peratures did not differ among the two reefs (F1,65=0.013,
P=0.910). However, on average LV–Lilipuna experienced
a 0.12°C lower daily minimum temperature during the
experimental period (F1,65=4.636, P=0.035) which led
to an overall greater daily temperature range (F1,65=5.250,
P=0.025) at LV–Lilipuna (1.27°Cday−1) compared to
HV–Reef 14 (1.16°Cday−1). Daily integrated light avail-
ability (Oct 2014–Dec 2014) was 10.6mol photonsday−1
and did not differ among the two study sites (F1,68=0.004,
P=0.949), but varied from 1.2 to 25.9mol photonsday−1
over this period.
Physiology responses
Corals appeared fully pigmented with polyps extended
for most of the experiment. However, after 6days corals
in the heated treatment began to show visible signs of pal-
ing. After 8days of treatment exposure Fo was significantly
higher in corals from LV–Lilipuna relative to HV–Reef
14 (F1,88=7.559, P=0.007) (Fig.4a), and Fo tended to
be higher in corals from heated treatments, although this
trend was not significant (F1,88=3.179, P=0.078). Con-
versely, Fm did not differ according to environmental history
(F1,88=1.955, P=0.166) but declined in corals from heated
treatments (F1,88=6.439, P=0.013) (Fig.4b). Fv/Fm was
higher in HV–Reef 14 corals (F1,87=5.723, P=0.019) and
declined in the heated treatments (F1,87=44.562, P<0.001)
(Fig.4c) (Table1). No environmental history×temperature
treatment interactions were observed for Fo, Fm, or Fv/Fm
(F1,87≥0.167, P≥0.178) (Table1).
Environmental history, temperature treatment, and their
interaction did not affect total protein biomass (F1,45≥0.069,
P ≥ 0.519) or Symbiodinium densities (F1,44 ≥ 0.025,
P≥ 0.420) (Table 1) (Fig. 5a, b). Conversely, chloro-
phyll a concentrations (μgcm−2) were lowest in HV–Reef
14 corals relative to those from LV–Lilipuna (3.585 vs.
4.684μg cm−2) (F1,45 = 6.175, P= 0.017), and chloro-
phyll concentrations decreased in those corals exposed to
heated treatments (3.667 vs. 4.687μgcm−2) (F1,45=5.264,
P=0.026). However, no environmental history×tempera-
ture treatment interactions were detected (F1,45=0.024,

Marine Biology (2018) 165:56
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Page 7 of 15 56
P=0.877) (Fig.5c). The concentration of chlorophyll a
normalized to Symbiodinium cell (pg chlorophyll a cell−1)
was not affected by environmental history (F1,44=1.465
P = 0.233), temperature treatment (F1,44 = 1.006,
P=0.321), or their interaction (F1,44=0.547, P=0.464)
(Table1) (Fig.5d).
Antioxidant andimmunological responses
Superoxide dismutase activity (i.e., SOD) (F1,43=6.912,
P=0.012) and catalase activity (i.e., CAT) (F1,43=5.648,
P=0.022) differed according to environmental history. No
temperature treatment (F1,43≥2.537, P≥0.068) or envi-
ronmental history× temperature treatment interactions
were detected for SOD or CAT (F1,43≥0.007, P=0.920)
(Table2) (Fig.6a, b). Pooled across temperature treatments,
SOD and CAT activity, was 28 and 29% lower in LV–Lili-
puna corals relative to HV–Reef 14 corals (Fig.6a, b).
Both prophenoloxidase activity (i.e., PPO) (F1,43=5.447,
P= 0.024) and melanin synthesis activity (i.e., MEL)
(F1,43=9.054, P=0.004) differed according to environ-
mental history (F1,43=5.447, P=0.024) and temperature
treatments (F1,43≥4.879, P≤0.033), but not their inter-
action (F1,43≥0.771, P≥0.358) (Table2). PPO activity
was 33% lower in corals from LV–Lilipuna relative to those
at HV–Reef 14, and PPO declined by 48% in heated treat-
ments (Fig.6c). Environmental history and temperature
treatment effects on PPO were directly opposite to MEL,
where LV–Lilipuna corals exhibited 56% greater MEL activ-
ity relative to corals from HV–Reef 14, and MEL activity
increased by 37% in response to heating (Fig.6d).
Discussion
Thermal stress and bleaching can suppress coral immunity
(Couch etal. 2008), leaving corals vulnerable to opportun-
istic infections and disease (Miller etal. 2009). The initial
ability of corals to avoid thermal stress and to resist patho-
genesis (i.e., constitutive antioxidative and immune activity)
is an important driver of coral fate during environmental
perturbation (Mydlarz etal. 2010; Palmer and Traylor-
Knowles 2012). Therefore, the effect of environmental
history on coral antioxidant profiles/immune activity has
important implications for organismal performance, disease
susceptibility, and responses to local and climate stressors
(Couch etal. 2008). Environmental history is an important
factor influencing the response of corals to physiological
stress (Brown etal. 2000, 2002a; Ainsworth etal. 2016) and
the capacity of corals to acclimatize and/or adapt to climate
change (Palumbi etal. 2014; Dixon etal. 2015; Torda etal.
2017). As such, studying the ability of coral populations and
Symbiodinium (Mayfield etal. 2012) to tolerate tempera-
ture variability (Maynard etal. 2008; Barshis etal. 2010),
persistent high pCO2 (Fabricius etal. 2011), and variable
pCO2 (Kenkel etal. 2017) is critical to understanding the
500
700
900
a
1400
1600
1800
2000
F
m
b
0.5
0.6
0.7
Ambient Heated
F
v
/ F
m
c
F
o
LV–Lilipuna
HV–Reef 14
Temperature Treatment
†
*
*
†
Fig. 4 a Fo (minimum fluorescence yield), b Fm (maximum fluores-
cence yield), and c Fv/Fm (maximum photochemical efficiency) of
dark-adapted Symbiodinium photosystem II reaction centers in the
coral Montipora capitata from two Kāne‘ohe Bay reefs (LV–Lilipuna
vs. HV–Reef 14) exposed to two temperature treatments (Ambient vs.
Heated). Values represent mean±SE (n=20–25), and symbols indi-
cate significant effects (P<0.05) of temperature treatment (*) or reef
environmental history (†)

Marine Biology (2018) 165:56
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56 Page 8 of 15
biology of reef corals in a world of rapid and unprecedented
environmental change.
Montipora capitata from both reefs maintained their Sym-
biodinium densities, chlorophyll a per Symbiodinium cell,
and total protein content when exposed to short-term ther-
mal stress. However, chlorophyll a cm−2 and photochemical
performance (i.e., Fm and Fv/Fm) declined in corals from
both reefs when exposed to the heated treatment. The dis-
crepancy in heating effects reducing chlorophyll a cm−2 but
not affecting symbiont densities or chlorophyll a cell−1 may
in part be explained by methodology (i.e., sample variability,
statistical power), in addition to biological processes (i.e.,
photoacclimation), differences in tissue and skeletal opti-
cal properties (Wangpraseurt etal. 2012), and the internal
light environments where Symbiodinium reside. Fv/Fm dif-
fered according to reef environmental history, with higher
Fv/Fm at HV–Reef 14 under ambient and heated conditions,
suggesting environmental history affected properties of Sym-
biodinium photomachinery and rates of electron transport
(Warner etal. 2010). The heated treatment reduced Fm and
Fv/Fm, and indicates temperature-mediated damage to the
photosynthetic machinery (Lesser 1997; Jones etal. 1998;
Warner etal. 1999) and/or the activation of photoprotective
mechanisms (Hoegh-Guldberg and Jones 1999; Osmond
etal. 1999). The decreased photochemical performance,
along with visible tissue paling and reduced chlorophyll a
cm−2, confirms M. capitata were experiencing stress com-
mensurate with the onset of bleaching prior to the appreci-
able loss of symbiont cells. Declines in Fv/Fm often precede
reductions in Symbiodinium or photopigment densities (Fitt
etal. 2001; Rodrigues and Grottoli 2007). Indeed, short-term
laboratory experiments have shown M. capitata maintains
high symbiont densities despite loss of pigmentation from
thermal (Rodrigues and Grottoli 2007) and ultraviolet (UV)
Table 1 Statistical analysis
of environmental history and
temperature treatment effects on
Symbiodinium and Montipora
capitata physiology
Env. history environmental history of low variable pCO2 (LV–Lilipuna) or high variable pCO2 (HV–Reef
14), Treatment ambient (24.5°C) or heated (30.5 °C), Fo (minimum fluorescence yield), Fm (maximum
fluorescence yield), and Fv/Fm (maximum photochemical efficiency)of dark-adapted Symbiodinium photo-
system II reaction centers, SS sum of squares, df degrees of freedom
Bold P values represent significant effects (P<0.05)
Dependent variable Effect SS df F P
FoEnv. history 0.247 1 7.559 0.007
Treatment 0.104 1 3.179 0.078
Env. history×treatment 0.060 1 1.842 0.178
Residual 2.879 88
FmEnv. history 0.137×1061 1.955 0.166
Treatment 0.451×1061 6.434 0.013
Env. history×treatment 0.789×1061 1.128 0.291
Residual 6.158×10688
Fv/FmEnv. history 0.014 1 5.723 0.019
Treatment 0.106 1 44.562 <0.001
Env. history×treatment 0.394×10−3 1 0.167 0.684
Residual 0.206 87
Protein cm−2 Env. history 0.001 1 0.069 0.794
Treatment 0.002 1 0.277 0.601
Env. history×treatment 0.003 1 0.422 0.519
Residual 0.352 45
Symbiodinium cm−2 Env. history 0.042×1011 1 0.025 0.875
Treatment 0.431×1011 1 0.259 0.613
Env. history×treatment 1.104×1011 1 0.663 0.420
Residual 73.270×1011 44
Chlorophyll a cm−2 Env. history 14.219 1 6.175 0.017
Treatment 12.121 1 5.264 0.026
Env. history×treatment 0.056 1 0.024 0.877
Residual 103.614 45
Chlorophyll a cell−1 Env. history 0.378 1 1.465 0.233
Treatment 0.260 1 1.006 0.321
Env. history×treatment 0.141 1 0.547 0.464
Residual 11.361 44

Marine Biology (2018) 165:56
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Page 9 of 15 56
Fig. 5 Physiological responses
of Montipora capitata from two
Kāne‘ohe Bay reefs (LV–Lili-
puna vs. HV–Reef 14) exposed
to two temperature treatments
(Ambient vs. Heated). a Total
protein content, b Symbiod-
inium densities, c chlorophyll a
concentration per area of coral
tissue, and d chlorophyll a per
symbiont cell. Values represent
mean±SE (n=10–13), and
symbols indicate significant
effects (P<0.05) of tem-
perature treatment (*) or reef
environmental history (†)
0.6
a
0
1
2
b
0
2
4
6
8
c
0.4
0.2
Ambient
Temperature Treatment
Heated
LV–Lilipuna
HV–Reef 14
Protein (mg cm
-2
)
Chlorophyll a (µg cm
-2
)
Symbiodinium (106 cm
-2
)
*
†
0
2
4
6
d
Ambient Heated
Chlorophyll a (pg cell
-1
)
Table 2 Statistical analysis
environmental history and
temperature treatment effects
on antioxidant enzymes and
immune activity of Montipora
capitata
Env. history environmental history of low variable pCO2 (LV–Lilipuna) or high variable pCO2 (HV–Reef
14), Treatment ambient (24.5°C) or heated (30.5°C), SS sum of squares, df degrees of freedom
Bold P values represent significant effects (P<0.05)
Dependent variable Effect SS df F P
Superoxide dismutase (SOD) Env. history 0.776×1071 6.912 0.012
Treatment 0.393×1071 3.497 0.068
Env. history×treatment 0.001×1071 0.010 0.920
Residual 4.827×10743
Catalase (CAT) Env. history 39.576 1 5.648 0.022
Treatment 17.780 1 2.537 0.119
Env. history×treatment 0.045 1 0.007 0.936
Residual 301.310 43
Prophenoloxidase (PPO) Env. history 1.542 1 5.447 0.024
Treatment 3.908 1 13.800 <0.001
Env. history×treatment 0.243 1 0.859 0.359
Residual 12.176 43
Melanin (MEL) Env. history 4.804×1031 9.054 0.004
Treatment 2.589×1031 4.879 0.033
Env. history×treatment 0.409×1031 0.771 0.385
Residual 22.815×10343

Marine Biology (2018) 165:56
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56 Page 10 of 15
radiation stress (Grottoli-Everett and Kuffner 1995), suggest-
ing M. capitata and its Symbiodinium possess an especially
robust capacity for photoacclimation.
In all corals sampled, levels of the antioxidative
enzymes superoxide dismutase (i.e., SOD) and catalase
(i.e., CAT) were not significantly affected by tempera-
ture treatments. Rather, significant differences were only
observed as a function of reef environmental history. The
lack of change in antioxidative activity was an unexpected
observation, and we present three possible explanations
for this result: (1) constitutive levels of antioxidants pro-
vided adequate protection during short-term thermal chal-
lenge; (2) antioxidative responses are a secondary form
of defense not employed in the early stages of thermal
stress, as such enzymes are energetically costly to pro-
duce (Palmer etal. 2011a); or (3) other compounds such as
melanin (i.e., MEL) have dual function and exhibit some
antioxidant activity (Nappi and Christensen 2005) provid-
ing sufficient protection against cellular damage during the
onset of thermal stress. In other studies, the production of
specialized antioxidative enzymes (e.g., SOD and CAT)
is only induced after prolonged exposure (Downs etal.
2002). Melanisation may also function as a general accli-
matization response to environmental perturbation. In this
way, the melanin synthesis pathway may be an important
immune parameter activated in corals exposed to periodic
environmental stressors such as elevated irradiance and
temperatures. Regardless, the observation of site-specific
antioxidant profiles and melanin synthesis activity indicate
phenotypic differences in corals at these two locations,
potentially as a result of distinct pCO2 histories at these
locations. It is not known if the reef-specific differences
in coral phenotypes observed here reflect mechanisms of
acclimatization or local adaptation to a history of distinct
environmental conditions; however, such lines of inquiry
should be advanced to further our understanding of envi-
ronmental history effects on corals.
The melanin synthesis pathway was responsive to heated
treatments in corals from both reefs regardless of environ-
mental history. This pathway begins with the proteolytic
cleavage of inactive PPO to the active phenoloxidase (i.e.,
PO), and through a series of intermediate reactions ulti-
mately leads to the production and deposition of melanin
into coral tissues (Mydlarz and Palmer 2011; Nappi and
Christensen 2005). PPO levels typically drop upon induc-
tion of melanin production as reserves of the latent enzyme
are converted to their active form and consumed (Mydlarz
etal. 2008; Palmer etal. 2011a, b). This agrees with results
observed in the present study, where thermal stress caused
reductions in coral tissue PPO and simultaneous increases
in melanin production. However, other enzymes are also
capable of completing the melanin synthesis pathway. For
example, peroxidases can compete with PO for the hydroxy-
lation of tyrosine and subsequent melanin deposition (Nappi
Fig. 6 Immunological responses
of Montipora capitata from two
Kāne‘ohe Bay reefs (LV–Lilli-
puna vs. HV–Reef 14) exposed
to two temperature treatments
(Ambient vs. Heated). a Super-
oxide dismutase (SOD) concen-
tration, b catalase (CAT) activ-
ity, c prophenoloxidase (PPO)
activity, d melanin (MEL)
concentration. Values represent
mean±SE (n=11–12), and
symbols indicate significant
effects (P<0.05) of tem-
perature treatment (*) or reef
environmental history (†)
0
Δ Abs 490nm min
-1
mg prot
-1
a
50
150
250
Catalase
d
0
Superoxide dismutase
10
3
mg protein
-1
c
0
Melanin
Abs 490 nm mg tissu
e
-1
b
Prophenoloxidase
min
-1
mg prot
-1
*†
†
†
*
†
LV–Lilipuna
HV–Reef 14
Temperature Treatment
detaeHtneibmAdetaeHtneibmA
0.05
0.10
0.1
0.2
0.3
5
10
15

Marine Biology (2018) 165:56
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Page 11 of 15 56
and Vass 1993; Nappi and Christensen 2005), although these
enzymes were not measured here.
Melanin is a multifunctional compound that serves many
roles. It is important for both wound healing (Palmer etal.
2011b) and pathogen encapsulation (Ellner etal. 2007;
Mydlarz etal. 2008). It is also implicated in Symbiodin-
ium photoprotection (Palmer etal. 2010, 2011a), as it is a
known UV-absorbing molecule in mammals (Ortonne 2002;
Sugumaran 2002). Similarly, invertebrate photoprotection is
demonstrated in the water flea, Daphnia spp., where mela-
nisation is positively correlated to UV exposure (Rautio
and Korhola 2002). This photoprotective function was also
recently confirmed in sponges, where melanin produced
by symbiotic bacteria was protective against UV-induced
intracellular reactive oxygen species (Vijayan etal. 2017).
The sea fan, Gorgonia ventalina, displayed melanisation in
response to elevated temperatures (Mydlarz etal. 2008), and
higher constitutive levels of melanin and melanin-containing
granular cells have also been documented in coral species
considered resistant to thermal bleaching (Palmer etal.
2010). The exact role of coral melanisation in response to
increased temperature has yet to be elucidated, however, and
the causes and consequences of increased melanin synthesis
activity under short-term and prolonged thermal stress have
interesting implications for cellular adaptive mechanisms.
In the present study, we found that corals exhibit consti-
tutive differences in photobiology, chlorophyll a, antioxida-
tive enzymes, and immunity. However, environmental his-
tory effects did not interact with temperature treatments to
alter thermal stress response trajectories. Therefore, while
environmental history can shape the response of corals to
bleaching stress (Brown etal. 2002a), the specific environ-
mental conditions at the two reefs in the present study did
not influence the biological response of corals to short-term
thermal stress. Other physical or biological factors in addi-
tion to pCO2 history may also be responsible for influencing
the reef-specific responses observed here. Such factors may
include pathogen infections or immune response elicitors
(Palmer etal. 2011a) and their present and historical distri-
bution within Kāne‘ohe Bay (Aeby etal. 2010), as well as
low coral/high bare substrate cover and dissolved inorganic
nitrogen concentrations (Couch etal. 2008). However, such
factors do not appear to have played a significant role in the
present study. First, previous exposure to disease and physi-
ological stress can elevate coral immune activity (Mydlarz
etal. 2009; Palmer etal. 2011a), and historically, coral dis-
ease (i.e., Montipora white syndrome) prevalence is greater
in southern Kāne‘ohe Bay reefs proximate to LV–Lilipuna,
relative to central and northern reefs (Aeby etal. 2010).
However, we observed greater antioxidative (SOD and
CAT) and immune (PPO) activity at HV–Reef 14 in cen-
tral Kāne‘ohe Bay. Therefore, historical disease prevalence
does not explain greater antioxidant or immune activity at
HV–Reef 14. Alternatively, it is possible immune activity
in LV–Lilipuna corals surviving historically high disease
pressure (southern Kāne‘ohe Bay) is a consequence of resist-
ance/immunity to immune activity elicitors. Second, coral
cover at the two reefs are comparable (ca. 75%) and inor-
ganic nutrients within Kāne‘ohe Bay are not different from
those measured on offshore reefs (Cox etal. 2006), suggest-
ing the influence of coral cover and dissolved nutrients in
explaining differences among corals in the present study may
be minimal. Seawater temperature (Coles and Jokiel 1978)
and flow speed influence coral performance (Dennison and
Barnes 1988), and it is possible slightly cooler daily mini-
mum temperature at LV–Lilipuna (0.12°C) or other proper-
ties of seawater associated with residence time/flow (Lowe
etal. 2009) exerted influence here. In addition, differences in
holobiont traits due to seasonality (Fitt etal. 2000) or symbi-
ont abundance (Cunning and Baker 2014) at the time when
stress is applied can influence stress outcomes, and winter-
acclimation may have attenuated heating effects in corals in
the present study. Therefore, while pCO2 history remains the
most salient difference between LV–Lilipuna and HV–Reef
14 (Drupp etal. 2011, 2013) best explaining the distinct
responses of corals to short-term heating, the influence of
other physical factors should not be wholly dismissed.
Differences in symbiont communities among M. capitata
colonies (Stat etal. 2011) can also influence physiological
responses and stress outcomes (Sampayo etal. 2008; Cun-
ning etal. 2016). Montipora capitata in the Main Hawaiian
Islands are known to associate with both clade C and/or D
Symbiodinium (Stat etal. 2013), namely C31 and D1-4-6 (S.
glynnii) (Cunning etal. 2016; Wham etal. 2017). The latter
are often found in corals from reefs with a history of ther-
mal stress and/or variance and degraded water quality, such
as Kāne‘ohe Bay (Stat etal. 2013, 2015). Thus, the reef-
specific effects reported here may result from a combination
of several non-mutually exclusive factors including environ-
mental history (Brown etal. 2002a), host genotypes (Barshis
etal. 2010; Bongaerts etal. 2010), symbiont community
(Sampayo etal. 2008), and microbial consortia (Morrow
etal. 2015), as well as unidentified genetic mechanisms (i.e.,
gene expression plasticity, DNA methylation) (Kenkel and
Matz 2016; Putnam etal. 2016).
The role of environmental history in shaping coral physi-
ology remains an important and burgeoning field of inquiry
(Brown etal. 2002a; Middlebrook et al. 2008; Kenkel
etal. 2013a, b; Ainsworth etal. 2016; Kenkel and Matz
2016), especially in the context of thermal and pCO2 stress
(Fabricius etal. 2011; Noonan and Fabricius 2016; Gib-
bin etal. 2017; Kenkel etal. 2017). Environmental history
and phenotypic plasticity are important considerations for
predictions in the biology, ecology, and evolution of marine
organisms (Gaylord etal. 2015; Torda etal. 2017). Here,
distinct environmental histories of pCO2 variability did not

Marine Biology (2018) 165:56
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56 Page 12 of 15
interact with thermal stress to shape the suite of host and
symbiont responses. Nevertheless, environmental history
exerted strong influence over coral and Symbiodinium at
both ambient and elevated temperatures, emphasizing dif-
ferences among local reef environments even at small spatial
scales are important in determining coral holobiont perfor-
mance under favorable and challenging conditions. Finally,
the melanin synthesis pathway was significantly upregulated
during the early stages of thermal stress, and provides fur-
ther evidence that melanisation is an important general stress
response in corals exposed to warming seawater preceding
the onset of symbiont losses.
Acknowledgements We thank Dr. Eric H. De Carlo and colleagues
with NOAA PMEL and CRIMP CO2 program for Kāne‘ohe Bay pCO2
data, and two reviewers for suggestions that improved the manuscript.
Biological collections were performed in accordance with the state
of Hawai‘i Department of Land and Natural Resources Division of
Aquatic Resources permitting guidelines. CBW was supported by an
Environmental Protection Agency (EPA) STAR Fellowship Assistance
Agreement (FP-91779401-1). The views expressed in this publication
have not been reviewed or endorsed by the EPA and are solely those of
the authors. HMP was supported by NSF OCE-PRF 1323822. LDM
was supported by NSF 1017458, and CAR was supported by LSAMP
Bridge to Doctorate program. This is HIMB contribution number 1722,
SOEST contribution number 10328.
Compliance with ethical standards
Conflict of interest The authors declare they have no conflict of inter-
est.
Ethical approval All applicable international, national, and/or institu-
tional guidelines for the care and use of animals were followed.
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