There is a repeatable succession in the development of biofouling communities on new surfaces (Watson & Barnes, 2004 ). Calcareous polychaetes are often amongst the most prominent early metazoan colonizers (Stark, 2008 ). However, the factors governing succession are complex, and the effects of changed conditions remain unknown. Here, we aimed to investigate the effects of lowered pH (7.7), compared to controls (ambient, pH 7.9), on both established community structure and on the development of communities on newly exposed surfaces in a flow‐through, pH controlled experimental aquarium system. We used the biofouling community from the Ria Formosa Lagoon in southern Portugal as our test system. We further aimed to investigate these effects over an extended 100‐day period that covered multiple generations of the commonest species.

Evaluations of impacts on communities and the identification of susceptible assemblages are crucial to predicting responses of and impacts on ecosystems. To date, such assessments are rare and outcomes usually unclear (Dijkstra et al ., 2011 ; Hale et al ., 2011 ). Encrusting biofouling communities are ideal test systems, as they include species with CaCO 3 exoskeletons through to those lacking hard structures. This community is important worldwide, being the main colonizers and transformers of new surfaces in shallow marine environments. They also have great economic importance. In 2008 alone, the costs of managing and preventing marine biofouling were estimated at $15 billion for desalination systems and power plants and $7 billion for shipping worldwide (Jackson, 2008 ). The major biofouling organisms are sessile encrusting groups, typically bryozoans, calcareous tube‐dwelling polychaetes, sponges, ascidians and hydrozoans. Within these groups, several taxa, including spirorbid polychaetes, celleporellid bryozoans and sea squirts of the genus Ascidia , are unusual in having extremely large or global ranges. Understanding how this community responds to altered environments, especially acidified conditions, is thus important both scientifically and economically.

Ocean acidification poses great potential threats to organisms and ecosystems (Doney et al ., 2009 ; Constable et al ., 2014 ). Negative impacts of acidified environments have been documented in several groups (Orr et al ., 2005 ; Dupont et al ., 2008 ; Byrne, 2011 ), with species whose calcium carbonate (CaCO 3 ) skeletons are large proportions of their total biomass expected to be more strongly affected (Royal Society, 2005 ), especially in early developmental stages (Dupont et al ., 2008 ); but data supporting either contention are equivocal (Ries et al ., 2009 ; Kroeker et al ., 2010 ). One problem is that much of our knowledge is based on single species studies, which may be useful for identifying underlying mechanisms, but tell us little about the effects of lowered seawater pH on communities (Hale et al ., 2011 ).

The ascidians and sponge were identified to the genus level using 18s barcoding. Primers 18S‐SSUA NSF4 5′‐CTGGTTGATYCTGCCAGT‐3′, 18S‐SSUA NSR581: 5′‐ATTACCGCGGCTGCTGGC‐3′ in a standard PCR mix (Biotaq, Bioline, UK) with the following PCR conditions 94 °C 30 s, 40 cycles of 94 °C 30 s, 55 °C 30 s, 72 °C 1 min and a final step of 72 °C for 5 min.

Plastic tiles were preserved in ethanol and used to investigate the detailed structure and appearance of the fouling spirorbids. Selected areas were cut from tiles and cleaned in an ultrasonic bath. Observations of gold‐coated samples were made using a Jeol 820 SEM at 20 kV (Welwyn Garden City, UK).

Seawater was collected from each experimental tank with a clean 20 cm 3 plastic pipette and placed in a clean glass pyrex bottle (WB40/80; SciLabware Ltd, Stoke‐on‐Trent, UK). Saturated mercuric chloride in deionized water was added to seawater samples to a concentration of 0.05% when bottles were sealed with a ground glass stopper coated with a thin layer of ultrahigh vacuum grease (Apiezon; SPI supplies, West Chester, PA, USA) to block air exchange. Samples were then stored at 4 °C until analysis. Both TA and DIC were measured by the Plymouth Marine Laboratory as previously described (Findlay et al ., 2013 ). TA was measured in duplicate for each sample and the estimate of measurement error = 0.4%. Dissolved inorganic carbon was measured using a DIC analyser (Model AS‐C3; Apollo SciTech, Bogart, GA, USA). A measurement volume of 0.75 cm 3 was used, with up to five measurements per sample. Values outside a 0.1% range were excluded from the final result. Duplicate measurements provided an estimate of measurement error = 0.2%. Chemistry parameters were evaluated using the CO2SYS spreadsheet ( http://cdiac.ornl.gov/ftp/co2sys/CO2SYS_calc_XLS_v2.1/ ; Table 1 ).

The following were measured: temperature (°C), salinity (ppm), plus total phosphate (μmol kg −1 seawater), total silicates (μmol kg −1 seawater), total alkalinity (TA: μmol kg −1 seawater), total carbon dioxide (DIC: μmol kg −1 seawater) (Table 1 ). Seawater quality was assessed weekly using commercial Aquarium test kits. Using this system, ammonia, nitrite and nitrates were maintained well below 0.4, 0.2 and 5 mg L −1 , respectively.

Precolonized HDPE pipe and new surfaces of HDPE pipe, glass fibre tank walls and limestone tiles were all numbered to facilitate matching during the experiment. Photographs were taken of all substrata at the start and end of trials using a NIKON D80 (Tokyo, Japan) with NIKON DX SWMED IF Aspherical AF‐S NIKKOR 18–70 mm 1 : 3 5.5–4.5 GED lens. In precolonized trials, pipes were photographed and three sections analysed per tank (nine sections per pH treatment). For each section, spirorbids were counted in 8.25 cm 2 areas ( n = 10) and other taxa in 25 cm 2 areas because of the difference in density between taxa. Values were then recalculated and expressed as numbers 10 cm −2 . In all trials, there were zero values in some counts made, and data were not normally distributed even after log, double log or arcsin transformation. Data were therefore analysed using nonparametric Kruskal–Wallis tests with Bonferroni P value corrections when multiple tests were run.

The experimental circuit was fed with the microalgae Isochrysis galbana (clone T‐ISO, at 18 000 cells cm −3 per experiment), supplied in continuous flow to each tank by a peristaltic pump (ISMATEC, Wertheim, Germany). Chlorophyll‐a concentration was measured in each tank in vivo , using a portable fluorometer (10AU‐Turner Designs, Sunnyvale, CA, USA).

The flow‐through holding system consisted of six independently supplied and operated tanks all at 23 °C, with 3 at control pH (7.9) and 3 low pH (7.7), that is three independent replicates, for each pH treatment. Sea water was supplied from the Ria Formosa lagoon via a sand filter that removed all particles larger than 1.2 mm diameter, and performed partial removal down to 0.6 mm. This size range is significantly larger than the minimum dimensions of most polychaete and ascidian larvae (Stanwell‐Smith et al ., 1997 ). Each experimental tank was aerated and received 150 cm 3 min −1 of seawater from the header tank, maintained at 23 °C using an aquarium heater (NEWATT; Aquarium systems, Sarrebourg, France) equipped with a thermostat (±0.1 °C). Excess water overflowed, and the water in each tank was totally exchanged 3–4 times per day. Experimental and header tank temperatures were logged every 30 min (±0.1 °C, probe = ACQ210N‐TL; Aquatronica, Reggio Emilia, Italy). Seawater pH was continuously logged (ACQ210N‐PH; Aquatronica), and pH in experimental tanks was automatically controlled by CO 2 injection into the tank aeration supply. Injected CO 2 was controlled by an Aqua Medic pH Computer Set and solenoid valve. Experimental tanks were illuminated with daylight fluorescent lamps with a 12‐/12‐h light/dark regime. Conditions in the experimental system were stabilized for 1 month prior to initiation of experimental trials. CO 2 and temperature were monitored and controlled in real time. Salinity was measured with a VWR EC300 conductivity meter (Carnaxide, Portugal), and pH was also measured daily with an OxyGuard Handy pH meter (Farum, Denmark).

Experiments were performed in the experimental station of the Centre of Marine Science in the Ria Formosa lagoon, Portugal. The facilities are licensed for animal experimentation, and the experiments were covered by a Group‐1 licence (Direcção‐Geral de Veterinária, Portugal).

Results

In a pilot study of the effects of predicted change on the biofouling community at the CCMAR (Centre of Marine Science) experimental station (Ria Formosa lagoon, Portugal, 36°59′33″N 7°54′17″W), there was no temperature effect, but reduced pH affected both community structure and composition, when held at typical summer (24 °C) and autumn/spring (19 °C) values (online supporting material). In this investigation, we thus concentrated on acidification effects and conducted experiments at pH 7.9 (ambient) and pH 7.7 at a constant 23 °C and ambient salinity (Table 1). The pH reduction (0.2 pH units) was less than that predicted by the IPCC ‘business‐as‐usual’ scenario of a reduction of 0.3–0.4 pH units in oceanic surface waters by the year 2100, but will likely be achieved between 2055 and 2070.

On the precolonized substrata, the initial community was dominated by the spirorbid polychaete Neodexiospira pseudocorrugata which accounted for 79.5–92.6% of the individuals present (Fig. 1). The other species present in high enough numbers to analyse effects of altered conditions were ascidians from species of the genus Aplidium (0.8–1.1%) and Molgula (2.3–12.8%), plus a sponge of the genus Leucosolenia (4.3–6.5%). In the controls (pH 7.9), there were no differences between the start and end of the trials in numbers of spirorbids (H = 3.27, 1 df, n = 60, P = 0.07), sponges (H = 3.35, 1 df, n = 90, P = 0.07) and the ascidian Aplidium sp. (H = 0.01, 1 df, n = 108, P = 0.92). There was a small (29%), but significant, increase in the ascidian Molgula sp. numbers at the end (H = 12.31, 1 df, n = 99, P < 0.01). Conversely, after 100 days exposure to pH 7.7, even though at this lower pH neither calcite nor aragonite was undersaturated, the community was changed markedly, with fewer spirorbids (47.4%), but more sponge colonies (Leucosolenia sp., 29%). In the ascidians, Molgula sp. were more common (23.4%) and Aplidium sp less at 0.2%. For all four taxa studied, new recruits were observed in all treatments. Spirorbid numbers decreased significantly from 11.1 ± 1.2 to 2.0 ± 1.2 individuals per 10 cm2 (H = 13.21, 1 df, n = 50, P < 0.0001); numbers of the ascidian (Molgula sp.) increased fourfold from the start to end of the trials (H = 9.73, 1 df, n = 90, P = 0.001); whilst the second, less abundant, ascidian (Aplidium sp.) decreased by an order of magnitude (H = 6.61, 1 df, n = 108, P = 0.01); and the sponge Leucosolenia sp. increased 2.5‐fold (H = 13.49, 1 df, n = 90, P < 0.0001).

Figure 1 Open in figure viewer PowerPoint Numbers of the main components of the biofouling community on precolonized HDPE pipe before and after exposure to either pH 7.9 (control) of pH 7.7 (predicted year 2100 level). Values are mean per 10 cm2 ± SE; significant differences shown in figure as: *P < 0.01, **P < 0.001, ***P < 0.0001.

The major change in community composition was due to the marked reductions in numbers of spirorbids at low pH, even though estimates were conservative. The counts quoted above only included living N. pseudocorrugata. Dead and destroyed individuals were visible from the remaining scar. The proportions of total numbers of spirorbids that were dead were not significantly different between treatments at the start of trials on precolonized pipe (pH 7.9, 19.7% (5.4% SE); pH 7.7, 12.6% (4.6% SE); t = 1.01, P = 0.332, 17 df). However, the proportion of dead individuals was significantly higher in the pH 7.7 treatment than in controls at the end of the experiment [pH 7.9, 23.3% (3.8% SE), pH 7.7, 72.8% (5.1% SE); t = 7.78, P < 0.0001, 17 df].

Biofouling communities colonize different materials with varying success. We therefore placed clean sections of high‐density polyethylene (HDPE) pipe that were open, having been cut lengthwise (Fig. 2), and clean limestone tiles into our system and also monitored colonization of cleaned PVC tank walls in the controls and pH 7.7 trials over the duration of the experiment. All three surfaces were open, and this avoided the possibility that metabolic effects due to enclosed areas could alter pH conditions. Densities of spirorbids differed markedly on the various surfaces at the end of the 100‐day trials (pH 8: H = 80.46, 2 df, n = 113, P < 0.0001; pH 7.7 H = 28.8, 2 df, n = 113, P < 0.0001) (Fig. 3). In pH 7.9, spirorbid colonization of tank walls was higher than tiles (H = 41.83, 1 df, n = 83, P < 0.0001) and HDPE pipe (H = 54.93, 1 df, n = 95, P < 0.0001) and pipe were higher than tiles (H = 15.71, 1 df, n = 48, P < 0.001). Reduced pH lowered spirorbid numbers on tank walls by nearly sixfold, on pipe by 3.5‐fold and on tiles by nearly fivefold, and all of these were significant (H = 74.33, 1 df, n = 130, P < 0.0001; H = 18.65, 1 df, n = 60, P < 0.0001; H = 25.13, 1 df, n = 36, P < 0.0001, respectively). Numbers of other taxa were too low to analyse after 100 days on new substrata.

Figure 2 Open in figure viewer PowerPoint Section of HDPE pipe used in colonization trials.

Figure 3 Open in figure viewer PowerPoint Colonization of new surfaces by the spirorbid Neodexiospira pseudocorrugata after 100 days exposure to either pH 7.9 (control) or pH 7.7. Values shown are means ± SE and presented as numbers per 10 cm2. All before and after differences were significant (Kruskal–Wallis tests, H > 18.6 in all cases) at P < 0.0001, indicated on figure by ***.

Colonization of new surfaces by diatoms and filamentous algae was markedly different in the reduced pH trials compared to controls. It was not possible to quantify this effect from counts. Estimates were thus made visually from photographs of HDPE pipe surfaces and colonization levels classified into five categories from the lowest (1) to highest (5). Because this is a category analysis, nonparametric statistics were used to test for differences, and algal colonization in controls (pH 7.9; mean score = 4.4) was significantly higher than in low pH treatments (pH 7.7; mean score = 1.8; Mann–Whitney W = 45, P = 0.008, n = 11).

SEM analyses showed largely intact spirorbids with smooth outer surfaces from controls, but those at low pH were frequently ‘breached’ revealing internal structures (Fig. 4). XRD and SEM analyses confirmed the mineralogy as low magnesium calcite and an ultrastructure comprised predominantly of very small (<5 μm, Fig. 4) randomly aligned prismatic units, with little or no pitting or dissolution. There also appeared to be less binding matrix between prisms in spirorbid skeletons from the low pH treatments.