# Effect of ocean warming and acidification on a plankton community in the NW Mediterranean Sea

L. Maugendre, Jean-Pierre Gattuso, J. Louis, A. de Kluijver, S. Marro, K.

Soetaert, Frédéric Gazeau

► To cite this version:

L. Maugendre, Jean-Pierre Gattuso, J. Louis, A. de Kluijver, S. Marro, et al. Effect of ocean warming and acidification on a plankton community in the NW Mediterranean Sea. ICES Journal of Marine Science, 2015, 72 (6), pp.1744-1755. 10.1093/icesjms/fsu161. hal-03334805

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ICES Journal of Marine Science (2015), 72(6), 1744–1755. doi:10.1093/icesjms/fsu161

Original Article

Effect of ocean warming and acidification on a plankton
community in the NW Mediterranean Sea

L. Maugendre<fnref n="1" />,<fnref n="2" />*, J.-P. Gattuso<fnref n="1" />,<fnref n="2" />, J. Louis<fnref n="1" />,<fnref n="2" />, A. de Kluijver<fnref n="3" />, S. Marro<fnref n="1" />,<fnref n="2" />, K. Soetaert<fnref n="4" />, and F. Gazeau<fnref n="1" />,<fnref n="2" />

Maugendre, L., Gattuso, J.-P., Louis, J., de Kluijver, A., Marro, S., Soetaert, K., and Gazeau, F. Effect of ocean warming and acidification on a plankton community in the NW Mediterranean Sea. – ICES Journal of Marine Science, 72: 1744–1755.

Received 21 May 2014; revised 13 August 2014; accepted 18 August 2014; advance access publication 17 September 2014.

The effect of ocean warming and acidification was investigated on a natural plankton assemblage from an oligotrophic area, the bay of Villefranche (NW Mediterranean Sea). The assemblage was sampled in March 2012 and exposed to the following four treatments for 12 days: control (~360 µatm, 14°C), elevated pCO₂ (~610 µatm, 14°C), elevated temperature (~410 µatm, 17°C), and elevated pCO₂ and temperature (~690 µatm, 17°C). Nutrients were already depleted at the beginning of the experiment and the concentrations of chlorophyll a (chl a), heterotrophic prokaryotes and viruses decreased, under all treatments, throughout the experiment. There were no statistically significant effects of ocean warming and acidification, whether in isolation or combined, on the concentrations of nutrients, particulate organic matter, chl a and most of the photosynthetic pigments. Furthermore, ¹³C labelling showed that the carbon transfer rates from ¹³C-sodium bicarbonate into particulate organic carbon were not affected by seawater warming nor acidification. Rates of gross primary production followed the general decreasing trend of chl a concentrations and were significantly higher under elevated temperature, an effect exacerbated when combined to elevated pCO₂ level. In contrast to the other algal groups, the picophytoplankton population (cyanobacteria, mostly *Synechococcus*) increased throughout the experiment and was more abundant in the warmer treatment though to a lesser extent when combined to high pCO₂ level. These results suggest that under nutrient-depleted conditions in the Mediterranean Sea, ocean acidification has a very limited impact on the plankton community and that small species will benefit from warming with a potential decrease of the export and energy transfer to higher trophic levels.

**Keywords:** climate change, ocean acidification, ocean warming, oligotrophic area, plankton community, primary production.

**Introduction**

Anthropogenic carbon dioxide (CO₂) emissions are responsible for an important increase in atmospheric CO₂ partial pressure (pCO₂). The consequences of CO₂ emissions are an increase of surface ocean temperature expected to rise by 2–4°C by the end of this century based on the current emission rates (IPCC, 2013). About 25% of anthropogenic CO₂ emissions are absorbed by the ocean (Le Quéré et al., 2013), generating profound modifications of the ocean carbonate chemistry and referred to as “ocean acidification”. The pH of the surface ocean has decreased by 0.1 units since the beginning of the industrial era and is projected to decrease by an extra 0.3–0.4 units by the end of the present century (Orr, 2011). Seawater

warming and acidification are expected to significantly affect the carbon cycle through the changes in the functioning of marine organisms and communities.

CO₂ fluxes between the atmosphere and the ocean are partly driven by biological activity. In the surface mixed layer, the balance between the autotrophic fixation of CO₂ by primary producers and the consumption/mineralization of organic matter by the whole plankton community is referred to as the net community production (NCP). A system is referred to as autotrophic when production exceeds consumption and heterotrophic when consumption is higher than production. Depending on the atmospheric pCO₂ and sea surface temperature, the surface ocean potentially acts

<footnote n="1">¹Sorbonne Universités, UPMC Univ Paris 06, UMR 7093, LOV, Observatoire Océanologique, F-06230, Villefranche/mer, France</footnote>

<footnote n="2">²CNRS, UMR 7093, LOV, Observatoire Océanologique, F-06230, Villefranche/mer, France</footnote>

<footnote n="3">³Deltares, Delft, The Netherlands</footnote>

<footnote n="4">⁴Department of Ecosystems Studies, Royal Netherlands Institute for Sea Research (NIOZ), Yerseke, The Netherlands</footnote>

<footnote>*Corresponding author: e-mail: laure.maugendre@gmail.com</footnote>
as a sink of CO₂ for the atmosphere (Gattuso et al., 1998) when the surface mixed layer exports organic matter to the deep ocean. Primary producers have then a key role on carbon cycle and climate regulation.

In many marine plants and algae, RuBisCO, a key enzyme involved in CO₂ fixation, is generally limited at environmental CO₂ concentrations. An increase in CO₂ could therefore enhance phytoplankton photosynthesis and growth (Riebesell et al., 2007). However, experimental studies reported contrasting effects of elevated CO₂ on photosynthesis: stimulating, neutral or even inhibitory effects were found (see review by Riebesell and Tortell, 2011 and references therein). Such different responses could be due to species-specific differences in the efficiency of carbon concentrating mechanisms (CCMs; e.g. Giordano et al., 2005). Depending on the efficiency of their CCM, some species will benefit from elevated CO₂ conditions while others will not, provoking changes in community composition. Shifts toward smaller (e.g. Yoshimura et al., 2010) or larger phytoplankton cells (Tortell et al., 2002) have been observed while no change in the taxonomic composition have been reported for communities acclimated to large seasonal pH changes (Nielsen et al., 2012). Changes in community composition could have consequences on ecological processes (such as modifications of energy transfer to higher trophic levels) and biogeochemical cycling (i.e. modifications of the export to the deep ocean; Riebesell et al., 2007). With respect to ecosystem function, several studies have shown enhanced carbon fixation and an increase in the organic carbon to nitrogen ratio (e.g. Riebesell et al., 2007), while others reported limited or no effect (Feng et al., 2009).

Projected warming is also expected to significantly affect marine organisms and communities. Culture experiments and *in situ* sampling have shown increased metabolic rates as temperature increases (e.g. Eppley, 1972; Regaudie-de-Gioux and Duarte, 2012). However, as thermal tolerance greatly differs between species, some will face conditions outside of their tolerance range and will be forced to move their ecological niches (Gao et al., 2012 and references therein). Several experiments have shown that warming could induce a shift towards smaller phytoplankton species (Sommer and Lengfellner, 2008) as well as a tighter coupling between phytoplankton and bacteria with possible consequences on remineralization and carbon export (Hoppe et al., 2008). Furthermore, phytoplankton exhibits higher nitrogen to phosphate requirements in warmer conditions (Toseland et al., 2013), which might also impact biogeochemistry.

In recent years, an experimental effort has been initiated to investigate the effect of both drivers at the community level. Elevated temperature combined or not with elevated pCO₂ has been shown to enhance photosynthetic rates (Hare et al., 2007; Feng et al., 2009) as well as enhanced dissolved organic carbon relative to particulate organic production (Kim et al., 2011). A recent study focusing on the short- (2 weeks) and long-term (1 year) response of a diatom community showed that elevated pCO₂ and temperature, whether combined or taken in isolation, had an effect on the community structure, with a stronger influence of warming which induces a loss in species richness (Tatters et al., 2013). Another study supported the predominant effect of warming compared with acidification on bacterial phylogenetic composition (Lindh et al., 2013).

Nutrient availability is suspected to also have strong effects on the community response to ocean warming and acidification (Hare et al., 2007) and the great majority of past experiments have been performed under nutrient replete conditions. However, a large part (>60%) of the open ocean is characterized by oligotrophic

conditions with very low nutrient concentrations and rates of primary production (Dodds and Cole, 2007). Although the metabolic status (auto- vs. heterotrophic) of these areas is still under debate (Duarte et al., 2013; Williams et al., 2013) oligotrophic provinces represent ~30% of global oceanic primary production (Longhurst et al., 1995). Therefore, changes in the community composition and functioning in these regions could lead to significant changes in the global oceanic CO₂ sink. The Mediterranean Sea is a largely enclosed sea, presenting trophic status varying from mesotrophic in the Northwestern region to extremely oligotrophic in the Eastern basin. Despite these environmental constraints, the Mediterranean Sea hosts from 4 to 18% of the Earth's marine biodiversity (Bianchi and Morri, 2000) with a high percentage of endemic species. There is a growing concern on the effects of climate change and ocean acidification in this area, although, to the best of our knowledge, no experiment on the effect of elevated temperature and pCO₂ on natural plankton communities have been conducted to date.

In the present study, a Mediterranean plankton community sampled in winter was exposed to elevated temperature and pCO₂ as projected for the end of the century (respectively, +3°C and ×2 pCO₂). During 12 days, experimental bottles were placed in a control and a temperature-regulated outdoor tank. Parameters and processes such as carbonate chemistry, nutrients, particulate organic matter, pigments, cells abundance, and primary production were monitored regularly. Stable carbon isotope tracers (¹³C) were also used to measure carbon fixation.

# Material and methods

## Experimental setup

A volume of 300 l of seawater was sampled in the bay of Villefranche (France; 43°40′N, 7°18′E) at 5 m on 14 March 2012. Pumping was performed by a trace-metal clean pump activated by pressurized air from a diving tank, preventing any damage on the organisms. Seawater was sieved onto a 200-µm mesh to remove large organisms.

In the laboratory, seawater was transferred to an acid-cleaned 300 l tank. Labelled ¹³C-sodium bicarbonate was added to a final concentration of 19 µmol l⁻¹ corresponding to 0.83% of total dissolved inorganic carbon (DIC) concentration and increasing ⁸¹C-DIC by 760‰. A first set of 4 l acid-cleaned polycarbonate (PC) bottles (n = 24) was filled and hermetically sealed. CO₂-saturated filtered seawater was gently mixed with the remaining water to reach a calculated pCO₂ of ~750 µatm. This elevated pCO₂ seawater was then distributed to 4 l acid-cleaned polycarbonate bottles (n = 24). Half of the ambient and elevated pCO₂ bottles were placed in a 2 m³ tank installed on the pier of the Laboratoire d'Océanographie de Villefranche with a continuous flow of *in situ* seawater (20 l min⁻¹). The other half was placed in another identical tank in which temperature was maintained at ca. 3°C above *in situ* temperature. The four treatments, including three replicates bottles per sampling day, were: Control (C; ambient pCO₂ and temperature), Ocean Warming (OW; ambient pCO₂ and elevated temperature), Ocean Acidification (OA; elevated pCO₂ and ambient temperature), and Greenhouse (G; elevated pCO₂ and temperature). The bottles were gently stirred every day to keep particles suspended. Light (natural sunlight) conditions were similar between the different treatments and were representative of surface conditions (~1 m depth). On several occasions, photosynthetic active radiation (PAR) was measured in the tanks with a spherical sensor
connected to an LICOR data logger, and daily (sunrise–sunset) averages ranged between 1025 and 1213 µmol photons m⁻² s⁻¹. Temperature was measured with a Seabird SBE37 temperature sensor and temperature in the elevated temperature tank was regulated using a COREMA® controller. Temperature regulation was performed throughout the experiment but, due to a technical problem, data were not logged on the first day.

## Sampling and analyses

After 2, 4, 8, and 12 days (thereafter referred to as d2, d4, d8, and d12), three bottles of each treatment were removed from the tanks. Samples for DIC were stored in 60 ml brown borosilicate bottles and poisoned with 10 µl saturated solution of mercuric chloride (HgCl₂). For total alkalinity (A_T) determination, 500 ml was filtered through 47 mm GF/F filter, poisoned with HgCl₂ and preserved at 4°C pending analysis. Aliquots of 20 ml for the determination of inorganic phosphate (PO₄³⁻), nitrate and nitrite (NOₓ = NO₃⁻ + NO₂⁻) were filtered through 0.2 µm polycarbonate filters cleaned with Merck Suprapur hydrochloric acid and rinsed with 18.2 MΩ cm⁻¹ ultra pure water. Samples were stored in 125 ml polyethylene (PE-HD) flasks, cleaned with Merck Suprapur hydrochloric acid, and acidified with Merck Ultrapur HCl, and finally processed using a spectrophotometric method with a Liquid Waveguide Capillary Cell optic fibre, allowing the determination of nanomolar concentrations phosphate and NOₓ with detection limits of 1 and 9 nmol l⁻¹, respectively, and a precision (coefficient of variation) of ~7% for both parameters (Adornato et al., 2007; Zimmer and Cutter, 2012). Samples for the determination of silicate were stored in acid-cleaned PE vials, poisoned with a saturated solution of HgCl₂ and kept at 4°C pending analysis using an AXFLOW AA3 auto-analyzer. A 2 ml aliquot of seawater was added to 80 µl of glutaraldehyde for the analysis of community composition. Samples were snap-frozen in liquid nitrogen and kept at -80°C pending analysis on a FACSCalibur flow cytometer. For pigment analyses, 0.5–1 l of seawater were filtered on 25 mm GF/F membranes which were stored at -80°C pending extraction and analysis on an Agilent Technologies 1200 series following the protocol of Ras et al. (2008). For particulate organic carbon (POC) concentration and its isotopic signature (δ¹³C-POC), 1 l of seawater was filtered through pre-combusted and preweighted 25 mm GF/F filters under low pressure. Filters were dried at 60°C for 24 h then stored at room temperature in the dark. For measurement of δ¹³C-DIC, samples (20 ml) were poisoned with 10 µl of HgCl₂ and stored at room temperature in the dark. The remaining seawater in each 4 l polycarbonate bottle was used for measurements of community metabolism (see below).

DIC was determined immediately after opening the bottles on triplicate 1.2 ml subsamples using an inorganic carbon analyser (AIRICA, Marianda, Kiel, Germany) coupled to an infrared gas analyser (LI-COR 6262). This instrument was calibrated before sample analysis against a certified reference material provided by A. Dickson (Scripps Institution of Oceanography, San Diego, CA, USA; batch 114). The average precision (SD) of all measurements (n = 52; ran in triplicates) was 0.7 µmol kg⁻¹. A_T was determined on triplicate 50 ml subsamples by potentiometric titration on a Metrohm Titrando 80 titrator coupled to a glass electrode (Metrohm, electrode plus) and a thermometer. The pH electrode was calibrated daily on the total scale using TRIS buffers of salinity 35 provided by A. Dickson. Measurements were carried out at 25°C and A_T was calculated as described by Dickson et al. (2007). Along the experiment, standards provided by A. Dickson (batch 108)

were used to check precision and accuracy (n = 18; 2.9 and 3.7 µmol kg⁻¹, respectively). The parameters of the carbonate system were determined from DIC, A_T, temperature, and salinity using the R package seacarb (Lavigne et al., 2014). To take into account the uncertainty of the measured input parameters during the calculation of the carbonate chemistry parameters, a Monte-Carlo procedure was applied. One thousand values were randomly chosen between the mean ± SD of each measured parameter and mean ± SD of each computed parameters is reported.

POC samples were analysed for organic carbon content and isotope ratios on an elemental analyser (Thermo Electron Flash EA 1112) coupled to a Delta V isotope ratio mass spectrometer (IRMS). For DIC isotope analyses, a 2 ml helium headspace was created in the vials and samples were acidified with 2 µl of phosphoric acid (H₃PO₄; 99%). After equilibration for 30 min, the CO₂ concentration and its isotopic composition in the headspace were measured on the EA-IRMS.

Data of the ¹³C-labelling study were expressed in the delta notation (δ) relative to Vienna Pee Dee Belemnite standard. The carbon isotope ratio was calculated as

$$R_{\text{sample}} = \left( \frac{\delta^{13}\text{C}_{\text{sample}}}{1000 + 1} \right) \times R_{\text{VPDB}}, \quad \text{with } R_{\text{VPDB}} = 0.0111797.$$

The ¹³C fraction was calculated as:

$$^{13}\text{F} = \frac{^{13}\text{C}}{^{13}\text{C} + ^{12}\text{C}} = \frac{R}{R + 1},$$

where R = ¹³C/¹²C.

The excess ¹³C was obtained as Δ¹³F = ¹³Fsample - ¹³Fbackground. Absolute incorporation rates were calculated as ¹³C-POC = Δ¹³F × [POC]sample (µmol C l⁻¹; De Kluijver et al., 2010).

Finally, ¹³C-concentrations were converted to total fresh POC:

$$\text{New - POC} = \left( \frac{\Delta^{13}\text{F}_{\text{POC}}}{\Delta^{13}\text{F}_{\text{DIC}}} \right) \times [\text{POC}],$$

where Δ¹³FPOC and Δ¹³FDIC are the excess values, [POC] is a concentration in µmol C l⁻¹.

NCP is then calculated as:

$$\text{NCP} - ^{13}\text{C} = \frac{\Delta\text{New - POC}}{\Delta t},$$

where ΔNew-POC is the differences between two consecutive sampling days.

## Primary production and community respiration

NCP (NCP-O₂) and community respiration (CR-O₂) were measured using the oxygen light–dark technique. Gross community production was measured using the ¹⁸O-labelling method (gross primary production, GPP-¹⁸O).

Before sunrise, three 60 ml biological oxygen demand bottles were sampled from each PC bottle. One bottle was immediately fixed with Winkler reagents to determine the initial O₂ concentration. A transparent and a dark bottle were incubated in the outdoor tanks for 24 h for estimating NCP-O₂ and CR-O₂, respectively. O₂ concentrations were measured using an automated Winkler titration technique with potentiometric endpoint detection. Analyses were performed with a Metrohm Titrando 888 with
a Metrohm ion electrode. Reagents and standardizations were similar to those described by Knap et al. (1996). NCP-O₂ and CR-O₂ were estimated by regressing O₂ values against time, and CR was expressed as negative values. GPP (GPP-O₂) was calculated as the difference between NCP-O₂ and CR-O₂. The combined errors were calculated as:

$$SE_{x-y} = \sqrt{(SE_x^2) + (SE_y^2)}$$

For the ¹⁸O-labelling technique, samples were transferred from each PC bottle to two 60 ml transparent glass bottles and sealed. One bottle was directly poisoned with 10 µl saturated solution of HgCl₂ to estimate the natural isotopic composition and the other bottle was spiked, with 50 µl of 97% H₂¹⁸O to reach a final isotopic composition δ¹⁸O-H₂O of 335‰. After 12 h incubations in the outdoor tank (from sunrise to sunset), samples were poisoned using HgCl₂ and stored upside down in the dark at room temperature pending analysis. Measurements were performed at KU Leuven (Belgium). A headspace of 3 ml was created with helium and allowed to equilibrate for 30 min (¹⁸O-O₂ measurements). The extracted water was injected into helium-flushed vials (¹⁸O-H₂O measurements). Pure CO₂ (100 µl) was then added and samples were allowed to equilibrate for 24 h. δ¹⁸O-H₂O was therefore measured as δ¹⁸O-CO₂. Determinations of δ¹⁸O-O₂ and δ¹⁸O-CO₂ were accomplished using an elemental analyzer (Flash HT/EA) coupled to a Delta V IRMS. An overflow technique was used to limit air contamination of the needle. For δ¹⁸O-O₂, the internal standard used to correct the data and survey instrumental deviation was air from the outside. For δ¹⁸O-CO₂, a calibration was performed with a VSMOW standard. GPP rates (µmol O₂ l⁻¹ d⁻¹) were calculated using the following equation (Kiddon et al., 1995):

$$GPP_{-18}O = \left[ \frac{\delta^{18}O-O_2_{final} - \delta^{18}O-O_2_{init}}{\delta^{18}O-H_2O - \delta^{18}O-O_2_{init}} \right] \times [O_2]_{init}$$

where δ¹⁸O-O₂ init and δ¹⁸O-O₂ final are measured δ¹⁸O-O₂ before and after incubation (‰), respectively, δ¹⁸O-H₂O is the final isotopic composition of the labelled water (‰), and [O₂] init is the O₂ concentration before incubations (µmol l⁻¹). The overall error was estimated using a Monte-Carlo procedure as described earlier.

## Statistics and data availability

Data are presented as averages ± SD (or ± SE for metabolic rates). Due to the small number of replicates (×3), PERMANOVA analyses were performed using the R package RVAideMemoire (Hervé, 2013) to test for differences in parameters/processes between the four different treatments. These analyses were performed considering two interacting factors (pCO₂ and temperature) and one blocking factor (time) over 1000 permutations and a significant effect was considered when *p* < 0.05. Cumulative metabolic rates were calculated for the whole experimental period. Values for days when no incubations were performed were obtained by linear interpolation and the cumulative values were then summed up for the experimental period. The data reported here as well as complementary parameters are freely available in Pangaea: http://doi.pangaea.de/10.1594/PANGAEA.834159

## Results

In the two tanks, temperature naturally varied by ~2°C between day and night (Figure 1). The natural average temperature was 14.3 ± 0.3°C while it was on average 17.2 ± 0.7°C in the elevated temperature tank (average difference: 2.8°C). In treatments C and OW, pCO₂ was on average 364 ± 14 µatm (pHₜ 8.12 ± 0.02) and 414 ± 12 µatm (pHₜ 8.07 ± 0.01), respectively. Elevated pCO₂ conditions were on average 613 ± 22 µatm (pHₜ 7.92 ± 0.01) and 690 ± 28 µatm (pHₜ 7.88 ± 0.02) for treatments OA and G, respectively. The targeted pCO₂ levels were not reached most likely as a consequence of significant outgassing while bottles were filled. Aₜ averaged for all treatments was 2568 ± 4 µmol kg⁻¹ (Table 1) and did not vary significantly between treatments and sampling days (Table 2).

All dissolved inorganic nutrients were close to the detection limit. The concentration of NOₓ and silicate did not vary between treatments (Table 2; Figure 2a and b) but the phosphate concentration was significantly lower under elevated pCO₂ (*F* = 13.19, *p* < 0.05; Table 2, Figure 2c). Phosphate was on average 13 ± 1 nmol P l⁻¹ and NOₓ (NO₃⁻ + NO₂⁻) concentrations remained constant (75 ± 20 nmol N l⁻¹), after an initial decrease between d0 and d2 (Figure 2b). Silicate concentrations did not vary with time and averaged 1.0 ± 0.1 µmol Si l⁻¹ (Figure 2a).

The concentration of POC did not significantly differ between treatments (Table 2) with an overall mean of 11 ± 1 µmol C l⁻¹ (Figure 3a). PON concentrations were low (mean: 0.8 ± 0.1 µmol N l⁻¹), except on d2 in the OW and OA treatments where concentrations reached 1.9 ± 0.4 µmol N l⁻¹ (Figure 3b). The particulate organic C : N ratio was high with a global average of 15 ± 1 (6 ± 1 for OW and OA on d2) and was not different between treatments (Table 2). As for POC and PON, the concentration of chlorophyll a (chl a) did not differ between treatments (Table 2) but varied significantly throughout the experiment (Figure 3c). It increased from an overall mean of 0.9 ± 0.1 µg l⁻¹ on d0 to 1.1 ± 0.0 µg l⁻¹ on d2. After d2, it decreased in all treatments to reach an average final concentration of 0.3 < 0.1 µg l⁻¹.

In terms of phytoplankton group succession during the experiment, diatoms (as represented by fucoxanthin pigments), prymnesiophytes (19'-hexanoyloxyfucoxanthin), and cyanobacteria (zeaxanthin) were detected in the samples with a large dominance

<figure><img src="image_2.png" /><figcaption>**Figure 1.** Temperature during the experiment. Blue: *in situ* temperature (Control and Ocean Acidification treatments). Red: *in situ* + 3°C (Ocean Warming and Greenhouse treatments).</figcaption></figure>
<table><caption>**Table 1.** Carbonate chemistry parameters in the control (C), ocean warming (OW), ocean acidification (OA), and greenhouse (G) treatments (average ± SD).</caption><thead><tr><th></th><th>Alkalinity<br>(µmol kg<sup>-1</sup>)</th><th>DIC<br>(µmol kg<sup>-1</sup>)</th><th>pCO<sub>2</sub><br>(µatm)</th><th>pH<sub>T</sub></th></tr></thead><tbody><tr><td colspan="5"><strong>Day 0</strong></td></tr><tr><td>C</td><td>2566 ± 1</td><td>2278 ± 0</td><td>366 ± 2</td><td>8.11 &lt; 0.01</td></tr><tr><td>OW</td><td>2567 ± 2</td><td>2286 ± 0</td><td>428 ± 1</td><td>8.05 &lt; 0.01</td></tr><tr><td>OA</td><td>2570 ± 2</td><td>2386 ± 0</td><td>622 ± 3</td><td>7.91 &lt; 0.01</td></tr><tr><td>G</td><td>2565 ± 1</td><td>2352 ± 0</td><td>618 ± 2</td><td>7.92 &lt; 0.01</td></tr><tr><td colspan="5"><strong>Day 2</strong></td></tr><tr><td>C</td><td>2566 ± 2</td><td>2283 ± 2</td><td>377 ± 2</td><td>8.10 &lt; 0.01</td></tr><tr><td>OW</td><td>2563 ± 0</td><td>2284 ± 2</td><td>431 ± 5</td><td>8.05 &lt; 0.01</td></tr><tr><td>OA</td><td>2565 ± 1</td><td>2371 ± 6</td><td>586 ± 18</td><td>7.94 ± 0.01</td></tr><tr><td>G</td><td>2566 ± 4</td><td>2374 ± 2</td><td>666 ± 2</td><td>7.89 &lt; 0.01</td></tr><tr><td colspan="5"><strong>Day 4</strong></td></tr><tr><td>C</td><td>2566 ± 2</td><td>2270 ± 3</td><td>356 ± 6</td><td>8.12 &lt; 0.01</td></tr><tr><td>OW</td><td>2566 ± 2</td><td>2273 ± 1</td><td>405 ± 1</td><td>8.07 &lt; 0.01</td></tr><tr><td>OA</td><td>2568 ± 2</td><td>2383 ± 3</td><td>620 ± 17</td><td>7.92 ± 0.01</td></tr><tr><td>G</td><td>2566 ± 3</td><td>2386 ± 3</td><td>716 ± 22</td><td>7.86 ± 0.01</td></tr><tr><td colspan="5"><strong>Day 8</strong></td></tr><tr><td>C</td><td>2568 ± 5</td><td>2277 ± 4</td><td>363 ± 12</td><td>8.12 ± 0.01</td></tr><tr><td>OW</td><td>2567 ± 1</td><td>2279 ± 1</td><td>415 ± 1</td><td>8.06 &lt; 0.01</td></tr><tr><td>OA</td><td>2567 ± 1</td><td>2385 ± 4</td><td>629 ± 13</td><td>7.91 &lt; 0.01</td></tr><tr><td>G</td><td>2573 ± 3</td><td>2380 ± 2</td><td>669 ± 14</td><td>7.89 &lt; 0.01</td></tr><tr><td colspan="5"><strong>Day 12</strong></td></tr><tr><td>C</td><td>2577 ± 11</td><td>2281 ± 6</td><td>359 ± 23</td><td>8.12 ± 0.02</td></tr><tr><td>OW</td><td>2569 ± 1</td><td>2275 ± 3</td><td>404 ± 5</td><td>8.07 &lt; 0.01</td></tr><tr><td>OA</td><td>2568 ± 0</td><td>2383 ± 2</td><td>617 ± 4</td><td>7.92 &lt; 0.1</td></tr><tr><td>G</td><td>2569 ± 5</td><td>2387 ± 8</td><td>708 ± 22</td><td>7.87 ± 0.01</td></tr><tr><td colspan="5"><strong>Average</strong></td></tr><tr><td>C</td><td>2569 ± 5</td><td>2278 ± 5</td><td>364 ± 14</td><td>8.12 ± 0.02</td></tr><tr><td>OW</td><td>2566 ± 2</td><td>2279 ± 6</td><td>414 ± 12</td><td>8.07 ± 0.01</td></tr><tr><td>OA</td><td>2568 ± 2</td><td>2382 ± 6</td><td>613 ± 22</td><td>7.92 ± 0.01</td></tr><tr><td>G</td><td>2568 ± 3</td><td>2377 ± 12</td><td>690 ± 28</td><td>7.88 ± 0.02</td></tr></tbody></table>

Total alkalinity ($A_T$) and dissolved inorganic carbon (DIC) were measured, while the partial pressure of CO$_2$ ($pCO_2$) and pH$_T$ were estimated based on DIC and $A_T$ using seacarb (see the Material and method for more details).

of prymnesiophytes (28% on average; Figure 4a-c). Variations in the concentration of diatoms and prymnesiophytes were similar to chl *a* variations, i.e. an increase during the first 2 days followed by a general decrease. Variations in zeaxanthin concentrations indicate that the abundance of cyanobacteria increased from d0 to d8 then declined at the end of the experiment. This is consistent with flow cytometer data which show the same dynamics for the cyanobacteria *Synechococcus* (Figure 4f) with a significant correlation between the two techniques (HPLC and flow cytometry; *r* = 0.86, *p* < 0.01, *n* = 52). The abundance of *Prochlorococcus*, another cyanobacteria, increased until d4 then decreased (data not shown). Pigments and flow cytometry data showed a specific response to elevated temperature alone for cyanobacteria (zeaxanthin: *F* = 6.98, *p* < 0.05; *Synechococcus* by flow cytometry: *F* = 6.11, *p* < 0.05; Table 2, Figure 4c and f). The abundance of pico-eukaryotes was significantly different at elevated $pCO_2$ (*F* = 8.69, *p* < 0.05; Table 2), a difference that could be attributed to a transient higher abundance of this group in the OA treatment on d2 (data not shown). The abundance of viruses and heterotrophic prokaryotes decreased with time (Figure 4d and e). There was no difference between treatments for heterotrophic prokaryotes (Table 2, Figure 4d) and a significant temperature effect was found for viruses (*F* = 5.57, *p* < 0.05; Table 2) due to two very high values

in the C and OA treatments on d8 (Figure 4e), this effect disappeared when these two values were omitted.

Based on the O$_2$ light-dark technique, no significant difference between treatments was observed for the considered metabolic processes (NCP-O$_2$, GPP-O$_2$, and CR-O$_2$; Table 2). NCP-O$_2$ ranged from -1.6 ± 0.8 to 2.8 ± 0.2 µmol O$_2$ l$^{-1}$ d$^{-1}$ while CR-O$_2$ ranged from -4.3 ± 0.8 to -0.8 ± 1.3 µmol O$_2$ l$^{-1}$ d$^{-1}$ (Figure 5a and b). GPP-O$_2$ varied from -0.15 ± 1.5 to 5.74 ± 3.1 µmol O$_2$ l$^{-1}$ d$^{-1}$ (Figure 5c). NCP-O$_2$ did not show a clear temporal trend except for G treatment for which NCP-O$_2$ decreased from autotrophic to heterotrophic conditions throughout the experiment and in general, very large variations were observed for all treatments (Figure 5a). GPP-$^{18}$O followed the decreasing trend of chl *a* (Figure 5d) and was significantly increased under elevated temperature (*F* = 15.82, *p* < 0.01; Table 2) with a significant interaction with $pCO_2$ (*F* = 7.28, *p* < 0.05). No significant correlation was found between GPP estimated by the two methods (GPP-$^{18}$O and GPP-O$_2$; *r* = 0.26, *p* > 0.05, *n* = 48). The cumulative GPP-$^{18}$O was 33.0 ± 3.4 and 29.0 ± 2.3 µmol O$_2$ l$^{-1}$ for C and OA treatments, respectively. OW and G treatments presented higher cumulative values with 34.8 ± 2.3 and 38.5 ± 2.4 µmol O$_2$ l$^{-1}$, respectively.

$\delta^{13}$C-DIC in natural sample was ~3‰ (data not shown) and reached, as expected, 759 ± 18‰ after addition of labelled $^{13}$C-sodium bicarbonate. $^{13}$C-DIC did not significantly change during the course of the experiment and did not differ between treatments (Figure 6a, Table 2). The transfer from labelled DIC to POC was very rapid and efficient, allowing the detection of $^{13}$C-POC enrichment on the first sampling day (d2; Figure 6b), and saturation was achieved already at d4. $^{13}$C-POC enrichment did not differ between treatments (Table 2). $\delta^{13}$C-POC increased to a final enrichment of 501 ± 23‰. NCP-$^{13}$C was lower than NCP-O$_2$ exhibiting a decreasing trend (Table 3) and did not significantly differ between treatments (Table 2). Cumulative NCP-$^{13}$C was 11.8 ± 0.6 and 11.4 ± 0.2 µmol C l$^{-1}$ for C and OA, respectively. The warmer treatments had slightly higher values of 12.3 ± 0.3 and 12.1 ± 0.5 µmol C l$^{-1}$ for OW and G, respectively.

Discussion

This experiment was designed to study the effects of ocean warming and acidification on the composition and functioning of an oligotrophic plankton community in the coastal NW Mediterranean Sea. The elevated temperature condition was very well controlled with an average offset between ambient and elevated temperature of 2.8 ± 0.4°C. The current rate of warming in the coastal NW Mediterranean Sea has been estimated to range from 0.026 to 0.033°C yr$^{-1}$ (Bensoussan et al., 2009), although temperature projections are difficult to obtain due to large regional differences. Given these rates, the average 2.8°C temperature increase as applied in our study was representative for the end of the century. Although $pCO_2$ was lower than targeted and therefore lower than commonly used in similar perturbation studies, high-$pCO_2$ values of ~610-690 µatm correspond to the level of atmospheric CO$_2$ projected for 2060 according to the RCP 8.5 scenario (Meinshausen et al., 2011).

The *in situ* sampling was performed 1 day after the maximum surface chl *a* concentration was measured in the Bay of Villefranche (data not shown; but see http://somlit-db.epoc.u-bordeaux1.fr/bdd.php). Consequently, nutrients were depleted with levels very close to detection limits. NO$_x$ was consumed rapidly in all treatments during the first 2 days of the experiment (from d0 to d2) and its
<table><caption>**Table 2.** Results of the permutational analysis of variance for selected parameters and processes.</caption><thead><tr><th rowspan="2"></th><th colspan="2">pCO<sub>2</sub></th><th colspan="2">temperature</th><th colspan="2">pCO<sub>2</sub>: temperature</th></tr><tr><th>F</th><th>p</th><th>F</th><th>p</th><th>F</th><th>p</th></tr></thead><tbody><tr><td>Carbonate chemistry</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Alkalinity</td><td>0.029</td><td>0.874</td><td>0.974</td><td>0.362</td><td>0.686</td><td>0.458</td></tr><tr><td>DIC</td><td>696.193</td><td>0.001*</td><td>0.171</td><td>0.685</td><td>0.656</td><td>0.443</td></tr><tr><td>pCO<sub>2</sub></td><td>555.319</td><td>0.001*</td><td>0.175</td><td>0.688</td><td>1.000</td><td>0.341</td></tr><tr><td>pH</td><td>678.988</td><td>0.001*</td><td>0.048</td><td>0.831</td><td>1.191</td><td>0.2917</td></tr><tr><td>Nutrients</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>NO<sub>x</sub></td><td>4.017</td><td>0.078</td><td>0.551</td><td>0.482</td><td>0.792</td><td>0.387</td></tr><tr><td>PO<sub>4</sub><sup>3-</sup></td><td>13.187</td><td>0.005*</td><td>0.038</td><td>0.836</td><td>2.589</td><td>0.152</td></tr><tr><td>Silicate</td><td>0.969</td><td>0.376</td><td>0.350</td><td>0.587</td><td>1.652</td><td>0.222</td></tr><tr><td>Particulate organic matter</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>POC</td><td>0.039</td><td>0.837</td><td>1.956</td><td>0.184</td><td>0.875</td><td>0.377</td></tr><tr><td>PON</td><td>0.000</td><td>0.990</td><td>0.135</td><td>0.729</td><td>3.240</td><td>0.093</td></tr><tr><td>POC : PON</td><td>0.050</td><td>0.855</td><td>0.030</td><td>0.876</td><td>3.713</td><td>0.079</td></tr><tr><td>Pigments</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Chl a</td><td>1.832</td><td>0.216</td><td>1.146</td><td>0.303</td><td>0.006</td><td>0.934</td></tr><tr><td>Fucoxanthin</td><td>1.254</td><td>0.285</td><td>0.548</td><td>0.479</td><td>0.004</td><td>0.966</td></tr><tr><td>19'-Hexafucoxanthin</td><td>0.384</td><td>0.537</td><td>2.570</td><td>0.125</td><td>0.210</td><td>0.656</td></tr><tr><td>Zeaxanthin</td><td>0.017</td><td>0.902</td><td>6.983</td><td>0.027*</td><td>0.037</td><td>0.842</td></tr><tr><td>Flow cytometry</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Synechococcus</td><td>0.332</td><td>0.566</td><td>6.106</td><td>0.028*</td><td>0.084</td><td>0.781</td></tr><tr><td>Pico-eukaryotes</td><td>8.694</td><td>0.016*</td><td>3.094</td><td>0.114</td><td>1.615</td><td>0.235</td></tr><tr><td>Nano-prokaryotes</td><td>0.057</td><td>0.816</td><td>0.093</td><td>0.788</td><td>0.042</td><td>0.837</td></tr><tr><td>Heterotrophic prokaryotes</td><td>0.145</td><td>0.729</td><td>1.392</td><td>0.268</td><td>0.002</td><td>0.974</td></tr><tr><td>Viruses</td><td>0.656</td><td>0.465</td><td>5.571</td><td>0.040*</td><td>0.291</td><td>0.588</td></tr><tr><td><sup>13</sup>C labelling</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td><sup>13</sup>C-DIC</td><td>1.746</td><td>0.204</td><td>0.214</td><td>0.642</td><td>4.592</td><td>0.057</td></tr><tr><td><sup>13</sup>C-POC</td><td>2.124</td><td>0.165</td><td>2.105</td><td>0.052</td><td>0.120</td><td>0.738</td></tr><tr><td>Metabolic rates</td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>GPP-<sup>18</sup>O</td><td>0.003</td><td>0.956</td><td>15.824</td><td>0.006*</td><td>7.283</td><td>0.021*</td></tr><tr><td>GPP-O<sub>2</sub></td><td>0.218</td><td>0.642</td><td>1.257</td><td>0.324</td><td>0.179</td><td>0.668</td></tr><tr><td>NCP-O<sub>2</sub></td><td>0.074</td><td>0.776</td><td>769.0</td><td>0.769</td><td>0.539</td><td>0.483</td></tr><tr><td>CR-O<sub>2</sub></td><td>0.139</td><td>0.728</td><td>1.683</td><td>0.240</td><td>0.035</td><td>0.849</td></tr><tr><td>NCP-<sup>13</sup>C</td><td>0.867</td><td>0.368</td><td>2.526</td><td>0.155</td><td>0.020</td><td>0.901</td></tr></tbody></table>

The "*" indicate significant effect (p < 0.05).

concentration significantly decreased while chl *a* concentration increased rapidly to a maximum on d2. This important chl *a* increase after enclosure, followed by decline owing to the depletion of inorganic nutrient, was also reported in other bottle experiments (De Madariaga and Fernandez, 1990; Scarratt et al., 2006) and can be due to a containment effect. In addition, it must be stressed that sieving onto 200 µm most likely limited the abundance of large zooplankton releasing predation pressure during the first days but favouring growth of micro-heterotrophs that in turn grazed on small phytoplankton. Although degradation pigments representative for grazer faecal pellets (phaeophytin *a* and phaeophorbid *a*) were close or even below detection limit throughout the experiment (data not shown), suggesting that there were no or very few zooplanktonic organisms, as the available seawater volume for sampling was not sufficient, zooplankton abundances were not estimated during this study. Based on pigment concentrations, prymnesiophytes were the dominant species and all phytoplankton groups decreased in abundance along the experiment, except for cyanobacteria. The dominance of small phytoplankton and the general decrease in biomass are consistent with post-bloom, nutrient-depleted conditions. Cyanobacteria and prymnesiophytes present a higher surface vs. volume ratio and have a higher affinity for nutrients than larger phytoplankton, cyanobacteria being even more competitive and able to grow under these very low nutrient conditions (e.g. Eppley

et al., 1969; Irwin et al., 2006). The dominance of pico- and nano-planktonic species at the end of the bloom period has already been observed in the Bay of Villefranche (Sheldon et al., 1992). While chl *a* concentrations showed very clear variations during the experiment, it was not the case for POC. These values presented large variations in between replicates and could explain the fact that we did not observe any temporal variation in POC.

The rates of community metabolism are comparable with those measured during a previous experiment performed in the bay of Villefranche in March 2003 (González et al., 2008). However, in contrast to this study, GPP-<sup>18</sup>O and GPP-O<sub>2</sub> were not correlated during our experiment. This can be explained by the relatively small range of values measured during our study in contrast to the large range of values (different seasons, depth, and sites) presented by González et al. (2008). Due to the limited amount of water available in the 4 l bottles, NCP-O<sub>2</sub> and CR-O<sub>2</sub> rates have been measured without replication. This led to relatively large uncertainties in the determinations of these rates and, even more important, in the estimates of GPP-O<sub>2</sub>. Although the same number of samples were available to estimate GPP-<sup>18</sup>O rates, the errors associated with this method were much smaller than with the O<sub>2</sub> light–dark techniques as GPP-O<sub>2</sub> is estimated based on two values (NCP- and CR-O<sub>2</sub>) associated to large uncertainties. Therefore, the <sup>18</sup>O-labelling approach appears much more reliable for estimating GPP rates than the classical
<figure><img src="image_3.png" /><figcaption>**Figure 2.** Concentration of inorganic nutrients as a function of time: (a) silicate, (b) NOₓ (NO₃⁻ + NO₂⁻), and (c) phosphate. Control (C), ocean warming (OW), ocean acidification (OA), and greenhouse (G) treatments. Symbols are for the three replicates of each treatment. Lines: solid (C), dashed (OA), dotted (OW), and dotted-dashed (G).</figcaption></figure>

<figure><img src="image_4.png" /><figcaption>**Figure 3.** Particulate organic matter and chl *a* as a function of time: (a) POC, (b) particulate organic nitrogen (PON), and (c) chl *a* measured by HPLC. Control (C), ocean warming (OW), ocean acidification (OA), and greenhouse (G) treatments. Symbols are for the three replicates of each treatment. Lines: solid (C), dashed (OA), dotted (OW), and dotted-dashed (G).</figcaption></figure>

light–dark technique when rates are low and only a small sample volume is available. However, it must be stressed that the ¹⁸O-labelling techniques does not allow estimating CR, which is critical for determining the autotrophic vs. heterotrophic behaviour of the community. Nevertheless, the observed decrease in GPP-¹⁸O during the experiment was consistent with the decrease in the phytoplankton biomass indicated by the pigments analyses.

The ¹³C enrichment was successful and showed decreasing primary production rates along the experiment which is consistent with the observed decrease in phytoplankton biomass and GPP

measured by the ¹⁸O method. However, the NCP estimated based on the ¹³C method (NCP-¹³C), representing the freshly labelled material produced by phytoplankton, was lower than NCP estimated by the light–dark method (NCP-O₂). The NCP-¹³C and NCP-O₂ were measured from incubation in different volumes (60 ml vs. 4 l) and over different periods (24 h vs. 2 or more days), which might explain part of this difference. Moreover, it is likely that, a significant part of the organic material produced was released in the DOC pool (Wood and Van Valen, 1990; Lopez-Sandoval *et al.*, 2011) which was not measured during our experiment.
<figure><img src="image_5.png" /><figcaption>**Figure 4.** Pigments (left panels) and flow cytometer counts (right panels) as a function of time: (a) fucoxanthin (typically diatoms), (b) 19'-hexanoyloxyfucoxanthin (19'-hex; prymnesiophytes), (c) zeaxanthin (cyanobacteria), (d) abundance of heterotrophic prokaryotes, (e) abundance of viruses, and (f) abundance of *Synechococcus*. Control (C), ocean warming (OW), ocean acidification (OA), and greenhouse (G) treatments. Symbols are for the three replicates of each treatment. Lines: solid (C), dashed (OA), dotted (OW), and dotted–dashed (G).</figcaption></figure>

During the recent years, an experimental effort to study ocean warming and acidification effects at the plankton community level has been initiated; however, there is still a clear lack of information for low productive oceanic regions (oligotrophic) that represent an important, and expanding surface of the global ocean (e.g. Irwin and Oliver, 2009), having then a significant role in carbon cycling. The present experiment was designed to investigate the effect of both ocean acidification and warming under conditions that prevail for

most of the year in the oligotrophic Mediterranean Sea, i.e. very low nutrient availability and a community dominated by small phytoplankton species.

In our study, no significant effects of elevated temperature and/or CO₂ were found for most parameters and processes (Table 2). As reported in other experiments (e.g. Feng et al., 2009), C:N ratio was not affected either by temperature or CO₂. The pCO₂ effect detected on the phosphorus concentration is most likely due to sampling
<figure><img src="image_6.png" /><figcaption>**Figure 5.** Community metabolism as a function of time: (a) NCP (NCP-O₂), (b) CR (CR-O₂), (c) gross primary production using the O₂ techniques (GPP-O₂), and (d) gross primary production using the ¹⁸O-labelling technique (GPP-¹⁸O). Control (C), ocean warming (OW), ocean acidification (OA), and greenhouse (G) treatments. Symbols are for the three replicates of each treatment. Lines: solid (C), dashed (OA), dotted (OW), and dotted-dashed (G).</figcaption></figure>

and/or analytical uncertainty. Indeed, during the whole experiment, concentrations varied within a very small range of ± 5 nmol l⁻¹ and for most of the sampling days, differences between replicates were larger than between treatments. The GPP-¹⁸O rates were higher in warmer treatment and even more enhanced when combined with high CO₂ conditions. This is in agreement with observed stimulations of the metabolism by temperature (Eppley, 1972; Toseland et al., 2013). Our data also support the previous findings of studies with higher nutrient levels in which elevated pCO₂ exerts a moderate effect on primary production rates unless it is combined with an increase in temperature (bottles incubation <3 l; Hare et al., 2007; Feng et al., 2009). An experiment performed in larger volumes (3000 l) has shown an increase in DOC production as well as photosynthetic activity and decrease in POC production in warmer and high CO₂ level conditions while gross community production remained unchanged (Kim et al., 2011, 2013).

It must be mentioned that although no study combining both drivers were conducted in oligotrophic conditions, ocean acidification alone has been reported to decrease DOC production in the

nutrient-depleted Okhotsk Sea at high CO₂ level while POC production was unchanged (Yoshimura et al., 2010). Furthermore, in iron-limited areas (Bering Sea and North Pacific) presenting similar chl a level than in our study, effects of ocean acidification were investigated and have shown different effect on POC and DOC accumulation that have been related to differences in community structure (Yoshimura et al., 2013). Indeed at the site where small phytoplankton species (70% picoaukaryotes and 20% *Synechococcus*) were dominant, no effect was detected, as in our study, while at the site where diatoms were dominating (75%), POC accumulation was smaller at high CO₂ levels. In our study, cyanobacteria (comprising *Synechococcus* and *Prochlorococcus* spp.) is the only taxonomic group that has shown enhanced abundance in warmer conditions though to a lesser extent when combined with high CO₂ level. Indeed, while the increasing tendency in both warming and greenhouse treatments was similar during the first sampling days, elevated pCO₂ appeared as unfavourable to this population after day 8 of our experiment. This is partially consistent with single-cell experiments that have shown no effect of increased CO₂ alone on *Synechococcus*
<figure><img src="image_7.png" /><figcaption>**Figure 6.** Concentrations of labelled (a) dissolved inorganic carbon (DIC) and (b) POC expressed in µmol C l⁻¹. Control (C), ocean warming (OW), ocean acidification (OA), and greenhouse (G) treatments. Symbols are for the three replicates of each treatment. Lines: solid (C), dashed (OA), dotted (OW), and dotted-dashed (G).</figcaption></figure>

<table><caption>**Table 3.** NCP based on ¹³C enrichment in the control (C), ocean warming (OW), ocean acidification (OA), and greenhouse (G) treatments (average ± SD).</caption><thead><tr><th rowspan="2">Day</th><th colspan="4">NCP-¹³C (µmol C l⁻¹ d⁻¹)</th></tr><tr><th>C</th><th>OW</th><th>OA</th><th>G</th></tr></thead><tbody><tr><td>2</td><td>1.93 ± 0.26</td><td>2.14 ± 0.05</td><td>1.69 ± 0.06</td><td>2.00 ± 0.06</td></tr><tr><td>4</td><td>1.52 ± 0.05</td><td>1.54 ± 0.13</td><td>1.60 ± 0.03</td><td>1.46 ± 0.13</td></tr><tr><td>8</td><td>0.76 ± 0.05</td><td>0.76 ± 0.02</td><td>0.71 ± 0.03</td><td>0.79 ± 0.02</td></tr><tr><td>12</td><td>0.52 ± 0.01</td><td>0.58 ± 0.03</td><td>0.51 ± 0.06</td><td>0.62 ± 0.08</td></tr></tbody></table>

growth rates and significant effects of temperature alone or in combination with acidification (greenhouse; Fu et al., 2007). However, in contrast to our results, growth rates were not significantly different between the elevated temperature and the greenhouse treatments. Furthermore, the same study reported no effect on *Prochlorococcus*, while increased abundances in the warmer treatment were observed in our study (data not shown). Finally, in a natural community of *Synechococcus* and *Prochlorococcus* spp. carbon fixation rates were also not affected by acidification under both nutrient replete and P-limited conditions (Lomas et al., 2012).

Extrapolating our findings to *in situ* oceanic conditions and facilitating potential future comparison studies necessitates a

discussion on the limits of the experimental design considered in the present study. Indeed, it must be stressed that our experiment was performed in relatively small-enclosed bottles (4 l) that undoubtedly led to some confinement effects which constrained the plankton community response (i.e. no turbulence, no nutrient re-supply, no movement through the euphotic zone, etc.; Scarratt et al., 2006). In our study, cyanobacteria species observed during the incubations were not N₂ fixers; however, it must be stressed that some strains have been shown to increase their N₂ fixation rates under high CO₂ conditions (e.g. Hutchins et al., 2009). The presence of N₂ fixers species in the community could have had influenced the results reported here. In addition, seawater was sieved through 200 µm to remove large organisms meaning that we did not take into account the complete community and therefore these findings are valid only when top-down control is negligible (Kim et al., 2013). Any potential change in the top-down pressure, due to change in macrozooplankton enhanced grazing in warmer and/or acidified condition, was therefore not investigated in this study. To assess the potential effect of climate change on communities of two or more trophic levels and trophic interactions (e.g. from phytoplankton to macrozooplankton), large mesocosm (~50 m³) studies are more appropriate as whole communities can be trapped and *in situ* temperature, irradiance and water masses are maintained close to “real world” conditions (Riebesell et al., 2010, 2013).

It must be stressed that a rise in primary production with elevated temperature is not foreseen on a global scale as many studies report a decrease of primary production as a consequence of a stronger stratification of the water column which limits nutrient supply to the surface mixed layer (Bopp et al., 2005). The observed increase in abundance of cyanobacteria in the warmer treatments in our study is in accordance with *in situ* observations, as it is recognized that these species have a wider temperature range than other phytoplankton species. Cyanobacteria will probably benefit from climate change as long as the temperature does not exceed their thermal tolerance (Morán et al., 2010; O'Neil et al., 2012). A potential shift toward small species could lead to diminished energy transfer to microzooplankton as their lipid content is much lower than it is for larger phytoplankton species (e.g. Von Elert and Wolff from, 2001). Furthermore, the carbon export efficiency depends on the community structure and a shift to smaller species will probably lead to a less efficient carbon export under elevated temperature (Bopp et al., 2005). As nutrients (N and P) were highly limiting, the absence of effect by ocean acidification alone does not appear as a surprise. Unfortunately, no data on dissolved organic nutrient concentrations are available and we could not precisely investigate any modifications in nutrient acquisition mechanisms (inorganic vs. organic) under the different treatments. Nevertheless, as no change of plankton community composition and functioning were shown at elevated pCO₂, it appears that the community did not take advantage of the increased CO₂ availability. To conclude, this study on the combined effect of both drivers has shown that ocean acidification and warming in isolation do not have the same effect on cells abundances and production rates than when combined, emphasizing the need to study these two drivers synergistically.

## Acknowledgements

Thanks are due to M. Houtekamer, P. van Rijswijk, and P. van Breugel of the Royal Netherlands Institute for Sea Research for carbon isotope measurements and access to their facilities, to S. Bouillon and Z. Kelemen at the KU Leuven for ¹⁸O determinations and access to
their laboratory, as well as to S. Alliouane for the preparation of the AIRICA and to C. Guieu for providing material. Thanks to E. Howes, A. Moya, A. Sallon and P. Mahacek for their help during sampling and for technical assistance. We also thank two anonymous reviewers for their helpful and constructive comments. This work was funded by the EC FP7 project 'Mediterranean Sea Acidification in a changing climate' (MedSeA; grant agreement 265103).

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