Increases in atmospheric carbon dioxide are leading to physical changes in marine environments including parallel decreases in ocean pH and increases in seawater temperature. This study examined the impacts of a six month exposure to combined decreased pH and increased temperature on the immune response and disease status in the blue mussel, Mytilus edulis L. Results provide the first confirmation that exposure to future acidification and warming conditions via aquarium-based simulation may have parallel implications for bivalve health. Collectively, the data suggests that temperature more than pH may be the key driver affecting immune response in M. edulis. Data also suggests that both increases in temperature and/or lowered pH conditions may lead to changes in parasite abundance and diversity, pathological conditions, and bacterial incidence in M. edulis. These results have implications for future management of shellfish under a predicted climate change scenario and future sustainability of shellfisheries. Examination of the combined effects of two stressors over an extended exposure period provides key preliminary data and thus, this work represents a unique and vital contribution to current research efforts towards a collective understanding of expected near-future impacts of climate change on marine environments.
Citation: Mackenzie CL, Lynch SA, Culloty SC, Malham SK (2014) Future Oceanic Warming and Acidification Alter Immune Response and Disease Status in a Commercial Shellfish Species, Mytilus edulis L. PLoS ONE 9(6): e99712. https://doi.org/10.1371/journal.pone.0099712
Editor: Pikul Jiravanichpaisal, Fish Vet Group, Thailand
Received: January 30, 2014; Accepted: May 16, 2014; Published: June 13, 2014
Copyright: © 2014 Mackenzie et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was funded by the project entitled: Shellfish Productivity in the Irish Sea (SUSFISH). SUSFISH is partially funded by the European Regional Development Fund (ERDF) through the Ireland Wales Territorial Co-operation (INTERREG 4A) Programme 2007-2013. INTERREG 4A is managed in Ireland by the Southern and Eastern Regional Assembly on behalf of the Irish Government, the Welsh Government and the European Commission (http://www.irelandwales.ie). The funders had no role in study design, data collection and analysis, decision to publish,or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Increases in atmospheric carbon dioxide are leading to physical changes in marine environments. Under the IPCC IS92a CO2 emission scenario, ocean pH is expected to decrease by 0.2–0.4 units by the year 2100 , . In addition to changes in ocean pH, organisms will also have to contend with simultaneous increases in seawater temperature with the current average sea surface temperature of 19.7°C predicted to rise to 22.7°C by the end of the century , .
The ability of marine organisms to adjust to future climate change conditions will be critical to their health and ultimate survival. Bivalves are unable to thermo-regulate and are considered poor regulators of haemolymph acid-base balance , . Consequently, pH and temperature changes in the environment will likely have direct influence on bivalve haemolymph pH and temperature. Additionally, any physiological mechanism associated with the bivalve circulatory system is liable to be affected under hypercapnic conditions. The bivalve immune system, for example, exists as an integrative part of the circulation system , . The circulating cells, or haemocytes, represent the major cellular component of the animal's immune response and are responsible for a number of defence activities including phagocytosis or encapsulation of foreign or diseased cells, release of reactive oxygen metabolites and enzymes, and secretion of cytotoxic molecules , , , .
While prior investigations of the consequences of near-future changes in ocean pH and temperature have demonstrated a number of effects on marine organism physiology including impacts on growth, calcification and acid-base status , –, less confirmed is how coinciding stressors might influence immune response in organisms . Changes to single environmental factors, particularly temperature – but also salinity , pH  and dissolved oxygen  have demonstrated impacts on various immunological parameters including haemocyte numbers, phagocytosis and oxidative burst response.
The immune system is a major physiological mechanism ensuring host survival in the battle with pathogenic or parasitic organisms . As stress impacts immune function and such mechanisms are the primary line of defence against pathogens, there is likely a strong link between immune response and the outbreak of disease in shellfish culture , . Thus, climate change impacts on immunological aspects of physiology may be paralleled by associated changes in bivalve disease status.
Global climate change has increased pathogen development and survival, disease transmission and host susceptibility in the last thirty years –. Increased disease development in certain ecosystems can contribute to species decline and even extinction. Disease dynamics in the marine environment are being influenced by physical, chemical and biological alterations driven by climate change and such impacts will affect the sustainability of the shellfish industry. Multiple climate drivers such as ocean acidification and warming ocean temperatures have the potential to promote pathogen range expansion, infected host invasions and native host decline ,  and tropical pathogen range contractions may also occur . Additionally, it is possible that certain pathogens will be detrimentally affected by climate change conditions via changes to their required conditions thus allowing a relatively small number of emergent pathogens to infect new hosts with little or no tolerance .
Previous studies have shown associations between parasites and changing ocean temperatures, with a warmer environment being more favourable to the parasite –. Higher temperatures resulting in lower oxygen levels may stress organisms thus increasing their susceptibility to disease , , . In the temperate zone, shorter milder winters are expected to increase the frequency and intensity of transmission of diseases . Certain parasites also have higher growth rates, higher reproductive ability and decreased generation time with higher temperatures , .
Prior investigation also indicates that ocean acidification and associated stressors will be negative for calcifying organisms with more energy being required to build shells, which will reduce the amount of energy available to find food, reproduce or resist parasites and disease , , , . It is possible that ocean acidification will also have a regulatory effect on certain parasite populations especially those with free-living developmental stages , . However it is also possible that increased pathogenicity will occur if certain parasites are less affected by an acidified environment than their respective hosts .
This research aimed to examine the impacts of exposure to future ocean warming and acidification on the immune response and disease status of a commercially important bivalve species, Mytilus edulis (Linnaeus). The impacts of six months exposure to lowered pH and increased temperature on immunological responses, including haemocyte counts, phagocytosis, and nitroblue tetrazolium (NBT) reduction, and the health status of the mussels by histological and molecular examination were investigated.
Materials and Methods
No specific permits were required for the study, which complied with all relevant regulations. The species collected in this study is not endangered or protected.
Mytilus edulis (mean shell length = 50.51 mm, SD = 3.68 mm) were collected from a sub-tidal population in the Menai Strait, North Wales, UK on May 3rd, 2011 and acclimatised in large (200 L) flow-through holding tanks (12.5°C±SD 0.26°C, pH 8.01±SD 0.08, Salinity 34± SD 1 psu, 12L:12D light regime) at the School of Ocean Sciences, Bangor for a 3 week time period. During this time, mussels were drip-fed concentrated algal feed (Instant Algae Shellfish Diet 1800, Reed Mariculture, Campbell, CA, USA; 40% Isochrysis sp., 15% Pavlova sp., 25% Tetraselmis sp. and 20% Thalassiosira weissglogii (Grunow); 52% protein, 16.1% lipid, 22.0% carbohydrate and 9.9% ash) at a ration of 27 mg dry mass mussel−1 day−1.
Experimental set-up and conditions for this study are described in detail in Mackenzie et al. . Briefly, M. edulis were exposed to current (∼400 µatm) and future (∼1000 µatm) pCO2 levels in an aquarium-based CO2 system. Current and future seawater conditions were simulated with seawater held at two temperatures (ambient, ambient +4°C) and two pH levels (ambient, ambient - 0.4 pH units) as predicted under the IPCC IS92a CO2 emission scenario . This set-up allowed for examination of the interactive effects of varying pH and temperature conditions and included four treatments: ambient pH at ambient temperature (ambient), ambient pH at ambient temperature +4°C (warming), reduced pH at ambient temperature (acidified) and reduced pH at ambient temperature +4°C (acidified+warming).
Following input of seawater from the Menai Strait to an internal settling tank, sea water was filtered and UV treated before delivery to each of four header tanks (150 L) representative of the four experimental treatments. In-line heaters (Elecro Titanium Digital Heater) and cooling units (Aqua Medic TITAN 200) were used to maintain experimental temperatures. A pH controller (Walchem dual input pH controller) regulated addition of CO2 (g) to achieve acidified treatments.
Experimental header tanks gravity-fed (flow rate ∼5 ml second−1) seawater to 20 replicate 1.5 L tanks per treatment with overflows running to waste. Eight individual mussels were assigned to each tank per treatment. Mussels were acclimated to temperature treatments at an increase of 0.5°C day−1. Mussel tanks were cleaned three times per week and mussels were fed according to amounts and rates described for the acclimatisation period.
Collection, storage and analyses of seawater carbonate samples are described in detail in Mackenzie et al. . Briefly, 60 mL total alkalinity and dissolved inorganic carbon (DIC) seawater samples were collected fortnightly and sent to the Carbonate System Facility (LIMS) at the National Oceanography Centre, Southampton for total alkalinity and DIC analysis. Temperature (Mettler Toledo SG2 SevenGO, MT Ltd., Leicester) and salinity (TMC V2 ATC) of samples were also recorded at time of collection. Additionally, 30 mL nutrient samples were collected, filtered and frozen for analysis of phosphate and silicate concentrations at the Scottish Marine Institute. All values were entered into the CO2SYS model  to determine seawater carbonate parameters (i.e. pH, pCO2, HCO3, CO3−2, ΩAr, ΩCa) using the thermodynamic constants of Mehrbach et al.  refitted by Dickson and Millero . Carbonate chemistry values for the four experimental treatments are provided in Table 1. Values represent the mean ±SD of bimonthly measures taken over the six months exposure period (n = 12 for each treatment).
An initial group of adult M. edulis (n = 5) were sampled to determine baseline immune status. Following six months of exposure, five mussels were randomly selected from five experimental tanks (i.e. replicates) within each treatment (n = 5). Haemolymph samples of 0.3–0.6 mL were withdrawn from the sinus of the posterior adductor muscle of each mussel using 2 mL syringes and 25 G ½ needles. Five mussels were bled consecutively so that total haemocyte counts, phagocytosis counts, and NBT analysis could be carried out concurrently.
Total haemocyte counts.
A 20 µL haemolymph subsample from each mussel was used to determine total haemocytes ml−1, counted using an improved Neubauer haemocytometer .
Phagocytosis assay was performed as previously described ,  with some modifications. Briefly, a 20 µL subsample of haemolymph was placed onto a glass slide and allowed to adhere for 20 min in a moist incubation chamber before the addition of 20 µL of fluorescein 5-isothiocyanate Isomer 1 (FITC) labelled E. coli (Sigma). Glass slides were then incubated for a further 20 min after which all were rinsed with Tris-HCl buffer solution containing 2% NaCl (pH 7.6). Unphagocytosed bacteria were counterstained with ethidium bromide (Sigma) and then rinsed clear with Tris-HCL. Triplicate counts of 200 cells were immediately carried out using a 488 nm emission filter on a Zeiss microscope.
Nitroblue tetrazolium (NBT) reduction.
A modified version of the nitroblue tetrazolium (NBT) assay, as described by Pipe  and Bussell et al.  was applied. Twenty µL of haemocyte suspension were added to 96 well plates (Fisherbrand) in triplicate for each sample. Negative and positive controls were incorporated. Following incubation for 20 min, 20 µL of nitroblue tetrazolium (NBT) (Sigma) (2 mg/ml in Tris–HCl buffer containing 2% NaCl (pH 7.6)) were added to one set of triplicate wells for each sample. Twenty µL Tris buffer were added to corresponding negative control wells. The well plates were then incubated for a further 20 min before centrifugation (1000 g at 10°C for 10 min). All wells were then washed with Tris-HCl buffer solution before another centrifugation (x2) (1000 g) after which 20 uL of methanol (50%) were added before a final centrifugation. The plates were air dried before the addition of 240 uL potassium hydroxide (KOH) and 280 uL dimethylsulphoxide (DMSO) to all plate wells. Optical density (OD) values were measured on a Dynex MRX-II spectrophotometer (λ = 620 nm).
An initial sample of adult M. edulis (n = 30) was screened from the Menai Strait in May 2011 to determine the health status of the mussels in the field. Following six months of exposure, twelve mussels were randomly selected from twelve experimental tanks (i.e. replicates) within each treatment for determination of health status by histological and molecular examination (n = 12).
DNA was extracted from the initial mussel sample (n = 30) (gill tissue stored in 95% ethanol) using 10% Chelex 100 resin , . DNA was extracted from the corresponding paraffin-embedded tissue of each mussel from the laboratory trial consisting of twelve mussels per treatment (n = 12). Deparaffinization of the tissue was carried out  and DNA extractions were undertaken using protein precipitate and cell lysis using the QIAamp DNA Mini Kit (Qiagen).
Standard polymerase chain reaction (PCR).
Several PCRs were carried out to determine (1) which Mytilus species was being screened, (2) general bacterial screening, (3) general microbial screening, (4) presence or absence of Marteilia refringens (Grizel et al) and general haplosporidian spp. screening.
(1) PCR for species identification: Initially a PCR was carried out to detect the nuclear DNA markers Me15/Me16  to confirm that M. edulis was being screened as hybrid zones consisting of M. edulis, Mytilus galloprovincialis (Lamarck) and hybrids of both parent species are known to occur on the southwest coast of the UK. The PCR mastermix was modified slightly to include 5× green buffer. Amplification was conducted in 25 µl of the reaction mixture containing 14 µl ddH2O, 5 µl 5× green buffer (Promega), 2.5 µl of each of the four deoxyribonucleotide triphosphates dNTPs (dATP, dCTP, dGTP, dTTP) (0.2 mM), 1.5 µl MgCl2 (25 mM stock), 0.5 µl of the primer Me15 (5′-CCAGTATACAAACCTGTGAAGA-3′), 0.5 µl of the primer Me16 (5′-TGTTGTCTTAATGGTTTGTAAGA-3′) (100 pmol/µL stock), 1 µl of Go Taq DNA polymerase (0.03 U) and 1 µl of total DNA. Negative controls consisting of ddH20 were used. The following conditions were used for the PCR in a thermocycler: 94°C for 30 s, 55°C for 45 s and 70°C for 90 s (40 cycles). The expected product size for M. galloprovincialis is 126 bp while in M. edulis it is 180 bp. In hybrids both bands occur simultaneously at 126 bp and 180 bp.
(2) Bacterial screening: PCR was carried out to screen for general bacterial species that might be present using the universal primers EUBB and EUBA , which amplify the entire 16srRNA region. The PCR mastermix was modified to include 5 µl 5× green buffer, 5 µl dNTP (0.2 mM), 0.5 µl of MgCl2 (25 mM stock), 0.25 µl of each primer (EUBB and EUBA) (100 pmol/µL stock) and 0.1 µl GoTaq DNA polymerase (0.03 U) per PCR reaction. The PCR reaction mix was made up to a volume of 20 µl using ddH20 and 5 µl of undiluted genomic DNA. The following conditions were used for the PCR in a thermocycler: 95°C for 5 min, 35 cycles of 95°C for 30 s, 55°C for 30 s and 72°C for 30 s with a final extension at 72°C for 10 min. The expected product size is 1.5 Kbp. A second pair of primers, UNIV16s EUB f933/UNIV16s EUB r 1387  which are specific for universally conserved bacterial 16s rDNA region were used for the bacterial screening. The PCR mastermix and thermocycling conditions were modified. The PCR mastermix was made up to a volume of 49 µl for each PCR reaction: 37.75 µl ddH20, 10 µl 5× green buffer, 1 µl dNTPs (0.2 mM), 3 µl MgCl2 (25 mM stock), 1 µl of each primer (100 pmol/µL stock) and 0.25 µl GoTaq DNA polymerase (0.03 U). 1 µl of undiluted genomic DNA was screened. The PCR was carried out in a thermocycler as follows: 95°C for 1 min, 35 cycles at 94°C for 20 s, 56°C for 30 s, 72°C for 30 s and a final extension at 72°C for 7 min.
(3) Microbial screening: Universal primers 18ScomF1/18ScomR1 , which amplify the 18Scom region, were used for a general health screen to assess if any other foreign DNA in mussel tissue could be detected.
(4) Screening for potential mussel parasites: A PCR protocol targeting the ITS1 was carried out to detect M. refringens . Several generic haplosporidian PCRs were carried out using the HAP-F1 and HAP-R3 primers  and the ssu980 and HAP-R1 primers , . Electrophoresis of the amplification products from all PCRs was conducted in a 2% agarose gel. Fifteen µL of ethidium bromide, EtBr, (10 mg/ml stock) was added to the agarose gel and the gel was run at 110 V for 60 min.
Direct sequencing was carried out on products amplified by eurofins MWG using ABI 3730xl 96-capillary DNA Analyzers. DNA from pooled PCR products (n = 3/n = 4) from multiple individuals in each sample were isolated and purified using the Qiaquick gel extraction kit (Qiagen) prior to direct sequencing Both the forward and reverse strands of DNA samples were sequenced commercially (MWG eurofins). Each sequence was matched against the National Centre for Biotechnology Information (NCBI) nucleotide database with BLASTn (Basic Local Alignment Search Tool) to identify the species present.
A transverse section of mussel tissue (∼1 cm2) containing mantle, gill, digestive gland and gonad tissue was excised and fixed in 96% ethanol initially and tissue sections were subsequently transferred to Davidson's fixative at 4°C for 48 hrs  before being processed. The fixed tissue was dehydrated through an ascending ethanol series and embedded in paraffin. 5 µm tissue sections were stained using haematoxylin and eosin. Tissue slides were scanned for any pathological or morphological observations and macro- or microparasites at a magnification of 40× and under oil at 100x. The location and intensity of each type of parasite were recorded and the prevalence and intensity of infection were calculated where possible.
All statistical analyses were carried out in SPSS 14.0 for Windows (2005). Two-way ANOVAs were applied to determine the effect of pH, temperature and their interaction on immunological parameters (p<0.05). Data was tested for normality and homogeneity of variances (Levene's Test). One-way ANOVA were used to directly compare treatments. If necessary, data was log-transformed to meet assumptions. Significant difference in the diversity and prevalence of each parasite group and pathological condition, with varying pH and temperature, was calculated using chi-square (χ2) analysis with significance determined at p<0.05.
Total haemocyte counts.
Two-way ANOVA identified a significant negative effect of increased temperature on haemocyte counts but neither an influence of pH nor interaction between pH and temperature (Table 2). Haemocyte counts varied significantly between groups (F = 24.336; p<0.001) with post-hoc tests revealing that baseline animals had significantly lower haemocyte counts than any experimental treatments (ambient (p<0.001), warming (p = 0.001), acidified (p<0.001), acidified+warming (p = 0.001)). Animals held at either ambient temperature treatment (ambient, acidified) had significantly higher haemocyte counts than those under increased temperature treatments at either pH (warming (p = 0.005, p<0.001, respectively), acidified+warming (p = 0.004, p<0.001, respectively)) (Fig. 1).
Baseline value shown in grey scale. Lowercase letters indicate significant differences.
Two-way ANOVA showed a significant positive effect of increased temperature on percentage of phagocytosed cells as well as a significant interaction between pH and temperature but no effect of pH (Table 2). The percentage of phagocytosed cells varied significantly between all groups (F = 3.846; p = 0.020) but failed assumptions of homogeneity of variances even after transformed. Post-hoc tests identified a significantly higher percentage of phagocytosed cells in haemolymph of mussels held at decreased pH and warming conditions (acidified+warming) than in baseline animals (p = 0.047) or at ambient temperature treatments at either pH (ambient (p = 0.016), acidified (p = 0.001)). Additionally, animals held at increased temperature and ambient pH (warming) had a significantly higher percentage of phagocytosed cell than those under ambient temperature and decreased pH (acidified) (p = 0.044) (Fig. 2).
Nitroblue tetrazolium (NBT) reduction.
Two-way ANOVA identified a significant positive effect of both decreased pH and increased temperature on NBT reduction but no interaction between the two factors (Table 2). NBT reduction varied significantly between treatments (F = 6.349; p = 0.002) with post-hoc tests revealing that mussels reared under ambient temperature and pH conditions (ambient) had significantly lowered reduction than all other experimental treatments (warming (p = 0.004), acidified (p = 0.022), acidified+warming (p<0.001)) Animals held under warming conditions at either pH (warming, acidified+warming) had significantly increased reduction than baseline animals (p = 0.012 and p = 0.002, respectively) (Fig. 3).
PCR and direct sequencing.
(1) A product of 180 bp was amplified in the Me15/Me16 PCR confirming the mussel species to be M. edulis (Table 3). Direct sequencing confirmed the mussel species to be M. edulis.
(2) In the PCR using the universal primers for bacteria screening, EUBB and EUBA , multiple products were amplified at 800 bp, 900 bp and 1000 bp. The second PCR used to screen for bacteria using the Univ16s EUBf and Univ16s EUBr primers amplified a product at ∼500 bp (Table 3). Several bacterial species were detected in the initial sample and in the mussels from all four treatments. The highest prevalence was observed in mussels from the initial sample (92%), ambient treatment (92%) and acidified+warming treatment (92%) followed by the acidified treatment (83%) and warming treatment (67%). In the bacterial screening, no DNA sequences were recovered for the products amplified in the EUBA and EUBB PCR. Five forward and five reverse sequences were obtained from the products amplified in the Univ16sEUBf and Univ16s EUBr PCR from a subsample of infected individuals (n = 6). Bacterial species detected included a Gammaproteobacterium sp. (99% Query coverage, 98% Maximum identity), Pseudomonas sp. (98% Query coverage, 95% Maximum identity), Pectobacterium carotovorium (Jones) (100% Query coverage, 97% Maximum identity), Xanthomonadaceae sp. (99% Query coverage, 98% Maximum identity) and Serratia sp. (99% Query coverage, 97% Maximum identity).
(3) A product (∼350 bp) was amplified in the 18Scom region using the Zhang et al.  modified PCR. This product was amplified in mussels from the initial sample, warming and ambient treatments but was absent in mussels from the acidified and acidified+warming treatments. The free-living amoebae Hartmannella vermiformis (Page) (85% Query coverage, 99% Maximum identity) was identified using the 18Scom PCR.
(4) No products were amplified in the ITS region using the M. refringens PCR (Table 3). A product was amplified (350 bp) using the generic haplosporidian primers  in eighteen individuals in the initial sample (60% prevalence), however, no products were amplified in mussels from any of the experimental treatments. Direct sequencing of the products to confirm haplosporidian identification in the initial sample using the generic haplosporidian primers was unsuccessful. Using the second pair of haplosporidian primers (ssu980 and HAP-R1), a product was amplified at 430 bp in a single individual in the initial sample, which also amplified a product using the HAP-F1 and HAP-R3 primers. No products were amplified in the mussels from the experimental treatments (Table 3). In the direct sequencing, two forward and two reverse DNA sequences were obtained from the replicate PCR products amplified by the single mussel in the ssu980 and Hap-R1 PCR , . Blastn analysis of the DNA isolated from M. edulis confirmed the DNA to be that from a previously undescribed haplosporidian (Accession no. KC852876 and KC852877) most similar to Minchinia chitonis (Lankester) (Accession no. AY449711.1, 95–100% Query coverage, 92–93% Maximum identity) and a haplosporidian of the Florida marsh clam Cyrenoida floridana (Dall) (Accession no. AY449712.1, 90–98% Query coverage, 93–95% Maximum identity). A phylogenetic tree was generated based on the majority rule jackknife consensus parsimony analysis with 1000 pseudo replicates with 75% character deletion using a heuristic search with 1000 random sequence additions each.
In total, seven parasite groups were detected in M. edulis. Haplosporidian like organisms (HLOs) were observed in the tissues of ten mussels (33% prevalence) from the initial sample (Table 4) but were not observed or detected in the final sample of mussels screened from any treatment group of the laboratory trial. The HLOs consisted of a cluster of cells of varying sizes (approximately 10 to 30 microns) each with an eccentric nucleus.
No single parasite species was detected across the initial sample and all four mussel treatment groups. However, the copepod Mytilicola intestinalis (Steuer) and the amoeba Hartmannella sp. were observed in the initial sample and three of the experimental treatment groups, being absent only in mussels from the warming treatment. The highest parasite diversity was observed in the initial sample (6 species) followed by the acidified treatment (4 species), the ambient and acidified+warming treatments (3 species) and warming treatment (2 species). A significant decrease in parasite diversity from the initial sample was observed in three of the experimental treatments (ambient (p = 0.046), warming (p = 0.014), acidified+warming (p = 0.046). All other diversity comparisons were not significant.
Most prevalence comparisons were significantly different for each parasite and treatment, with the parasite being either absent or the prevalence increasing or decreasing compared to the initial sample. However, some comparisons did have a similar prevalence. Comparing parasite prevalence within each treatment, in the initial sample most comparisons were significantly different except for ciliates and Nematopsis sp. (p = 1) and M. intestinalis and trematodes (p = 0.411). In the warming treatment, a similar prevalence was observed for ciliates and trematodes, the only parasite groups present (p = 1). Of the three parasite groups present in the ambient treatment, a significant difference in prevalence was observed between M. intestinalis and prokaryote inclusions (p = 0.0003) and Hartmannella sp. and prokaryote inclusions (p = 0.024). No significant difference was observed between M. intestinalis and Hartmannella sp. (p = 0.189). In the four parasite groups present in the acidified treatment mussels, most prevalence comparisons were significantly different (p<0.0001) except for M. intestinalis and Hartmannella sp. (p = 1). In the acidified+warming treatment, no significant difference (p = 1) was observed between trematodes and Hartmannella sp.
Ciliates were only present in mussels from the initial sample and the warming treatment, with prevalence quadrupled in the latter showing a significant difference in prevalence between both samples (p = 0.0002). An increase in the prevalence of M. intestinalis was observed in three of the four treatments (ambient, acidified, acidified+warming,) compared to the initial sample, with the prevalence increasing significantly (p = 0.026) in the acidified+warming treatment but was not significantly different in the ambient and acidified treatments (p = 0.3). A slight increase in the prevalence of trematodes from the initial sample was observed in the warming treatment, which was not significant (p = 0.6), whereas a significant decrease (p = 0.0056) in trematode prevalence was observed in the only other treatment (acidified+warming) in which trematodes were observed. Across all treatments, Nematopsis sp. was only present in one individual of the acidified treatment. Prokaryote inclusions were observed in a high proportion of mussels from the ambient and acidified treatments (58% and 83%, respectively), with a significant difference being observed between both treatments (p = 0.0001) (Table 4, Fig. 4).
In total, three pathological conditions were observed in the M. edulis. These conditions included melanin aggregates in the mantle tissue, lipofuscin within the kidney epithelium  and focal haemocyte infiltration in the vascular connective tissue (Table 5, Fig. 5). All three conditions were observed in the four treatments however, melanin aggregates were not observed in the initial sample nor haemocyte infiltration in the acidified treatment. Mussels with the highest prevalence of melanin aggregates and lipofuscin, within the kidney epithelium, were observed in the acidified+warming treatment while the highest prevalence of haemocyte infiltration was observed in the warming treatment. No significant difference was calculated for the number of pathological conditions observed between samples.
This is the first confirmation that coinciding exposure to future acidification and warming conditions has parallel implications for immune and disease status in M. edulis. Collectively, the data suggests that temperature more than pH may be the key driver affecting immune response in M. edulis. Haemocyte counts were lower under 16°C temperature conditions and phagocytosis activity increased under such conditions. Conversely, NBT reduction increased under 16°C conditions and/or pH 7.68 conditions, and therefore the effect of pH to immune function should not be discounted. Likewise, data suggests that both increases in temperature and/or lowered pH conditions may lead to changes in parasite diversity and prevalence, pathological conditions, and bacterial incidence in M. edulis. These changes may result in the removal or emergence of a parasite and an increase or decrease in parasite prevalence. No significant parasites such as M. refringens for example, were detected in this study, however, the detection of a previously undescribed HLO in the initial sample and its impact on that mussel stock, which has yet to be determined, is significant.
Previous research has shown that increased stress increases total haemocyte counts (THC) in bivalves , , . However, investigation into the specific effect of temperature stress on THC has demonstrated both decreases , ,  and increases  in marine bivalves. Our results suggest that temperature negatively influences THC as warming resulted in a decrease in circulating haemocytes.
Any metabolic adaptations to warming and/or acidification stressors could result in a transfer of energy resources to other physiological systems , potentially at the expense of maintaining haemocyte abundance. Therefore, the observed decline in THC may be indicative of an overall reduced energy state of M. edulis under stress conditions as been seen previously in marine bivalves under stress conditions . Similarly, such declines may be representative of an overall depletion of energy reserves due to the extended exposure period (which included the species' natural spawning windows). Any immunological response mounted by the species to deal with stressors may also incur energetic costs including altered energy metabolism, altered fatty acid composition and reduced reproductive investment  and thus, an animal that has to sustain such a response over time will inevitably incur reduced energy stores. These in turn may lead to a decreased ability to mount an effective immunological response particularly if parallel shifts in pathology and parasite loads coincide. Consistent increases in THC in all treatments from baseline values could be a result of either proliferation of cells, or transfer of cells from tissues into circulation  in response to stress conditions. Acidification conditions did not appear to affect THC. Examination of the sole effect of acidified seawater conditions by Bibby et al.  also found no effect on THC.
Phagocytosis is considered a good measure of immune competency in bivalves . Previous studies have demonstrated that phagocytic activity of haemocytes is affected by changes in both temperature  and acidification . Stress conditions predominantly cause decreases in phagocytic activity , , , , Conversely, the current study observed significant increased phagocytosis in M. edulis under increased temperature with potentially some effect of lowered pH. Such changes were paralleled by an increase in oxidative burst (NBT). Reduced oxidative burst has been shown to decline in other bivalve species under conditions of temperature stress . Increases in both immunological parameters may be indications of the general decline in condition of the animal and could also represent a higher “immunological load” as presented by changing pathogen and parasite condition induced by temperature and/or pH stressors.
A reduction in parasite diversity in the treatments compared to the initial sample may have been due to the artificial holding conditions in the laboratory or it may have been due to the altered pH, temperature and ecology of the parasite, such as the removal of other hosts. The HLOs observed in the initial sample were not observed in the mussels from the different treatments at the end of the trial. This is likely due to the removal of mortalities during the trial, which may have been been infected with the HLOs, while the survivors were uninfected giving a negative result in the final sample at the end of the trial.
Prokaryote inclusion bodies, which are bacterial cellular reserve material used during periods of special growth phases or under special environmental conditions , were not detected in the initial sample but were observed in both treatments held at 12°C and were most prevalent in the reduced pH treatment. A high prevalence of bacteria detected using PCR in all treatment samples would be expected due to the laboratory holding conditions.
The prevalence of M. intestinalis, which causes ‘red worm disease’, almost doubled in the acidified+warming treatment, when compared to the initial sample. Increasing temperature has been shown to be a factor influencing infestation rates , however, M. intestinalis was not detected in mussels from the warming treatment in this study but was present in all other three treatments which might indicate that this copepod prefers an environment with a reduced pH when seawater temperatures are elevated. Hartmannella sp., a free-living amoeba, was observed in most samples but was absent (warming) or significantly reduced (acidified+warming) in warming treatments indicating that this species may be sensitive to elevated seawater temperatures. Ciliates were only detected in the warming treatment but were absent in the acidified+warming treatment, which might indicate that they are sensitive to a reduced pH environment.
Trematodes were only observed in the two treatments held at 16°C, however, a prevalence reduction of almost a third was observed in the acidified-warming treatment. These results may indicate that the trematodes are sensitive to a more acidic and cooler environment, due to their absence in treatments held at 12°C.
In this study, all pathological conditions observed in the initial sample increased significantly in the four treatments, with an additional condition (melanin aggregates), which is part of a general inflammatory-like response in invertebrates , being observed in the laboratory trial mussels. Lipofuscin, lipid-containing residues of lysosomal digestion, is considered one of the ageing or “wear and tear” pigments and may be symptomatic of membrane damage or damage to mitochondria and is thus associated with the break down and absorption of damaged blood cells , . In this study, all mussel samples had lipofuscin within the kidney epithelium with the acidified+warming treatment having the highest prevalence. Haemocyte infiltration had a significantly higher prevalence in mussels in the warming treatment but was absent in mussels in the acidified treatment. A positive correlation between haemocyte number and increasing seawater temperature has been recorded in the Mediterranean mussel, M. galloprovincialis  and in clams, Ruditapes philippinarum (Adams & Reeve) . Monari et al.  suggested that the increased number of haemocytes found in the clam Chamelea gallina (Linnaeus) under elevated seawater temperatures, might have been a consequence of a mobilisation of cells from tissues to haemolymph, in order to respond to bacteria.
Intrinsic factors such as host immunity and pathogen virulence will be more difficult to predict in climate change studies . Knowledge of the rate of pathogen evolution and host evolutionary response is critical in predicting disease spread and subsequent effects on biodiversity , . Certain climate change driven stressors may also have a more negative impact on the parasite than on hosts . Consequently, it is apparent that climate change-disease interactions are complex , , , ,  and that an interdisciplinary approach should be taken .
Ocean warming and acidification pose substantial threat to marine bivalves and similar thermo-conforming species with limited ability to regulate haemolymph pH. The immune system of marine bivalves such as M. edulis may become comprised under such conditions. Furthermore, the reduced immune capacity of M. edulis to cope with climate-induced stressors such as ocean warming and acidification while dealing with co-occurring changes in parasite loads and pathogen incidence poses advanced threat to the health of the species. As such, parallel impacts to the immune status and pathology of M. edulis could have serious implications for global fisheries that subsist on the production and export of this commercially valuable species . In addition, any ecosystem services provided by M. edulis (e.g. uptake and recycling of energy and nutrients, bioturbation and bioirrigation of marine sediments, sediment/shoreline stabilization and habitat formation ) could also be threatened. We therefore highlight the need for further investigation into the long-term effects of collective climate-induced stressors to combined aspects of bivalve health and physiology so as to provide a comprehensive illustration of impacts, which could inform management and protection of M. edulis as well as other marine species of high global ecological and economic value.
Conceived and designed the experiments: CLM SAL SCC SKM. Performed the experiments: CLM SAL SCC SKM. Analyzed the data: CLM SAL. Contributed reagents/materials/analysis tools: SAL SCC SKM. Wrote the paper: CLM SAL SCC SKM. Maintenance of animals: CLM.
- 1. Bindoff NL, Willebrand J, Artale A, Cazenave A, Gregory J, et al. (2007) Observations: Oceanic Climate Change and Sea Level. In: Solomon S, Qin D, Manning M, Chen Z, Marquis M, Averyt KB, Tignor M, Miller HL, editors. Climate Change 2007: The Physical Science Basis. Contribution of Working Group I to the Fourth Assessment Report of the Intergovernmental Panel on Climate Change. Cambridge: Cambridge University Press. pp 385–482.
- 2. Nicholls RJ, Wong PP, Burkett VR, Codignotto JO, Hay JE, et al. (2007) Coastal systems and low-lying areas. In: Parry ML, Canziani OF, Palutikof JP, van der Linden PJ, Hanson CE, editors. Climate Change 2007: Impacts, Adaptation and Vulnerability.Contribution of Working Group II to the Fourth Assessment Report of the Intergovernmental Panel on Climate Change. Cambridge: Cambridge University Press. pp. 315–356.
- 3. Sabine CL, Feely RA, Gruber N, Key RM, Lee K, et al. (2004) The oceanic sink for anthropogenic CO2. Science (New York, NY) 305(367): 367–71.
- 4. Michaelidis B, Ouzounis C, Paleras A, Pörtner H (2005) Effects of long-term moderate hypercapnia on acid-base balance and growth rate in marine mussels Mytilus galloprovincialis. Mar Ecol Prog Ser 293: 109–118.
- 5. Whiteley NM (2011) Physiological and ecological responses of crustaceans to ocean acidification. Mar Ecol Prog Ser 430: 257–271.
- 6. Wootton EC, Dyrynda E, Ratcliffe N (2003) Bivalve immunity: comparisons between the marine mussel (Mytilus edulis), the edible cockle (Cerastoderma edule) and the razor-shell (Ensis siliqua). Fish Shellfish Immunol 15(3): 195–210.
- 7. Wootton EC, Pipe RK (2003) Structural and functional characterisation of the blood cells of the bivalve mollusc, Scrobicularia plana. Fish Shellfish Immunol 15(3): 249–262.
- 8. Pipe RK, Coles JA (1995) Environmental contaminants influencing immune function in marine bivalve molluscs. Fish Shellfish Immunol 5(8): 581–595.
- 9. Pipe RK, Farley SR, Coles JA (1997) The separation and characterisation of haemocytes from the mussel Mytilus edulis. Cell Tissue Res 289: 537–545.
- 10. Cao A, Novás A, Ramos-Martínez JI, Barcia R (2007) Seasonal variations in haemocyte response in the mussel Mytilus galloprovincialis Lmk. Aquaculture 263(1–4): 310–319.
- 11. Riebesell U, Zondervan I, Rost B, Tortell PD, Zeebe R, et al. (2000) Reduced calcification of marine plankton in response to increased atmospheric CO2. Nature 407: 364–367.
- 12. Portner HO, Langenbuch M, Reipschlger A (2004) Biological Impact of Elevated Ocean CO2 Concentrations: Lessons from Animal Physiology and Earth History. J Oceanogr 60(4): 705–718.
- 13. Berge JA, Bjerkeng B, Pettersen O, Schaanning MT, Øxnevad S (2006) Effects of increased sea water concentrations of CO2 on growth of the bivalve Mytilus edulis L. Chemosphere. 62(4): 681–7.
- 14. Gazeau F, Quiblier C, Jansen JM, Gattuso J-P, Middelburg JJ, et al. (2007) Impact of elevated CO2 on shellfish calcification. Geophys Res Lett 34: 1–5.
- 15. Miles H, Widdicombe S, Spicer JI, Hall-Spencer J (2007) Effects of anthropogenic seawater acidification on acid-base balance in the sea urchin Psammechinus miliaris. Mar Pollut Bull 54(1): 89–96.
- 16. Spicer JI, Raffo A, Widdicombe S (2007) Influence of CO2-related seawater acidification on extracellular acid-base balance in the velvet swimming crab, Necora puber. Mar Biol 151: 1117–25.
- 17. Wood HL, Spicer JI, Widdicombe S (2008) Ocean acidification may increase calcification rates, but at a cost. P Roy Soc B-Biol Sci 275(1644): 1767–73.
- 18. Gooding RA, Harley CDG, Tang E (2009) Elevated water temperature and carbon dioxide concentration increase the growth of a keystone echinoderm. Proc Natl Acad Sci USA 106: 9316–9321.
- 19. Callaway R, Shinn AP, Grenfell SE, Bron JE, Burnell G, et al. (2012) Review of climate change impacts on marine aquaculture in the UK and Ireland. Aquat Conserv 22: , 389–421.
- 20. Cheng W, Hsiao I-S, Hsu C-H, Chen J-C (2004) Change in water temperature on the immune response of Taiwan abalone Haliotis diversicolor supertexta and its susceptibility to Vibrio parahaemolyticus. Fish Shellfish Immunol 17(3): 235–43.
- 21. Parry HE, Pipe RK (2004) Interactive effects of temperature and copper on immunocompetence and disease susceptibility in mussels (Mytilus edulis). Aquat toxicol (Amsterdam, Netherlands) 69(4): 311–25.
- 22. Malagoli D, Casarini L, Sacchi S, Ottaviani E (2007) Stress and immune response in the mussel Mytilus galloprovincialis. Fish Shellfish Immunol 23(1): 171–7.
- 23. Monari M, Matozzo V, Foschi J, Cattani O, Serrazanetti GP, et al. (2007) Effects of high temperatures on functional responses of haemocytes in the clam Chamelea gallina. Fish Shellfish Immunol 22: 98–114.
- 24. Matozzo V, Marin MG (2011) Bivalve immune responses and climate changes: is there a relationship? Invertebrate Surviv J 8: 70–77.
- 25. Bussell JA, Gidman EA, Causton DR, Gwynn-Jones D, Malham SK, et al. (2008) Changes in the immune response and metabolic fingerprint of the mussel, Mytilus edulis (Linnaeus) in response to lowered salinity and physical stress. J Exp Mar Bio Ecol 358(1): 78–85.
- 26. Bibby R, Widdicombe S, Parry H, Spicer J, Pipe R (2008) Effects of ocean acidification on the immune response of the blue mussel Mytilus edulis. Aquat Biol 2: 67–74.
- 27. Cheng W, Li C-H, Chen J-C (2004) Effect of dissolved oxygen on the immune response of Haliotis diversicolor supertexta and its susceptibility to Vibrio parahaemolyticus. Aquaculture 232(1–4): 103–115.
- 28. Ellis RP, Parry H, Spicer JI, Hutchinson TH, Pipe RK, et al. (2011) Immunological function in marine invertebrates: responses to environmental perturbation. Fish Shellfish Immunol 30(6): 1209–1222.
- 29. Lacoste A, Malham SK, Gélébart F, Cueff A, Poulet SA (2002) Stress-induced immune changes in the oyster Crassostrea gigas. Dev Comp Immunol 26(1): 1–9.
- 30. Malham SK, Lacoste A, Gélébart F, Cueff A, Poulet SA (2003) Evidence for a direct link between stress and immunity in the mollusc Haliotis tuberculata. J Exp Zool 295A: 136–144.
- 31. Harvell CD, Mitchell CE, Ward JR, Altizer S, Dobson AP, et al. (2002) Climate warming and disease risks for terrestrial and marine biota. Science 296: 2158–2162.
- 32. Walther GR, Post E, Convey P, Menzel A, Permesan C, et al. (2002) Ecological responses to recent climate change. Nature 416: : 398–395. ISSN: 0028-0836.
- 33. Parmesan C, Yohe G (2003) A globally coherent fingerprint of climate change impacts across natural systems. Nature 421: 37–42. ISSN: 0028-0836.
- 34. Lafferty KD, Porter JW, Ford SE (2004) Are diseases increasing in the ocean? Ann Rev Ecol Evol S 35: 31–54.
- 35. Ward JR, Lafferty KD (2004) The elusive baseline of marine disease: are diseases in ocean ecosystems increasing? PLoS Biol 2: 542–547.
- 36. Thomas CD, Williams SE, Cameron A, Green RE, Bakkenes M, et al. (2004) Biodiversity conservation: Uncertainty in predictions of extinction risk/Effects of changes in climate and land use/Climate change and extinction risk (reply). Nature 430: 6995.
- 37. Epstein PR, Diaz HF, Elias S (1998) Biological and physical signs of climate change: focus on mosquito-borne disease. B Am Meteorol Soc 78: 409–417.
- 38. Patz JA, Campbell-Lendrum D, Holloway T, Foley JA (2005) Impact of regional climate change on human health. Nature 438: 310–317.
- 39. Jones KE, Patel NG, Levy MA, Storeygard A, Balk D (2008) Global Trends in Emerging Infectious Diseases. Nature 451: 990–93.
- 40. Pilling P, Hoffman I (2011) Climate change and animal genetic resources for food and agriculture - State of knowledge, risks and opportunities. FAO CGRFA Background Study Paper No. 53, p. 28.
- 41. Ford SE (1996) Range extension by the oyster parasite Perkinsus marinus into the northeastern United States: response to climate change? J Shellfish Res 15: 45–56.
- 42. Cook T, Folli M, Klinck J, Ford S, Miller J (1998) The relationship between increasing sea surface temperature and the northward spread of Perkinsus marinus (Dermo) disease epizootics in oysters. Estuar Coast Shelf Sc 40: 587–97.
- 43. Lafferty KD (2009) The ecology of climate change and infectious diseases. Ecology 90: 888–900.
- 44. Colwell RR (1996) Global climate and infectious disease: the cholera paradigm. Science 274: 2031–2025.
- 45. Harvell CD, Kim K, Burkholder JM, Colwell RR, Epstein PR, et al. (1999) Review: Marine ecology - Emerging marine diseases - Climate links and anthropogenic factors. Science 285: 1505–1510.
- 46. Pascual M, Rodo X, Ellner SP, Colwell R, Bouma MJ (2000) Cholera dynamics and the El Nino-Southern oscillation. Science 289: 1766–1769.
- 47. Harvell CD, Kim K, Quirolo C, Smith G, Weir J (2001) Mass mortality of Briareum asbestinum associated with the 1998 Caribbean coral bleaching. Hydrobiologia 460: 97–104.
- 48. Porter JW, Dustan P, Jaap WC, Patterson KL, Kosmynin V (2001) Patterns of spread of coral disease in the Florida Keys. Hydrobiologia 460: 1–24.
- 49. Harvell CD (2004) The rising tide of ocean diseases: unsolved problems and research priorities. Front Ecol Environ 2: 375–382.
- 50. Holmes JC (1996) Parasites as threats to biodiversity in shrinking ecosystems. Biodivers Conserv 5: 975–83.
- 51. Scott ME (1988) The impact of infection and disease on animal populations: implications for conservation biology. Conserv Biol 2: 40–56.
- 52. Kim K, Harvell CD, Kim PD, Smith GW, Merkel SM (2000) Fungal disease resistance of Caribbean sea fan corals (Gorgonia spp.). Mar Biol 136: 259–267.
- 53. Chubb JC (1979) Seasonal occurrence of helminths in freshwater fish. Part II. Trematoda. Adv Parasitol 17: 141–313.
- 54. Pojmanska T, Grabda-Kazubska B, Kazubski SL, Machalska J, Niewiadomska K (1980) Parasite fauna of five fish species from the Konin lakes complex, artificially heated with thermal effluents, and from Gopło Lake. Acta Parasitol Polonica 27: 319–357.
- 55. McDonald MR, McClintock JB, Amsler CD, Rittschof D, Angus RA, et al. (2009) Effects of ocean acidification over the life history of the barnacle Amphibalanus amphitrite. Mar Ecol Prog Ser 385: 179–187.
- 56. Kroeker KJ, Kordas RL, Crim RN, Singh GG (2010) Meta-analysis reveals negative yet variable effects of ocean acidification on marine organisms. Ecol Lett 13: 1419–1434.
- 57. Amaral V, Cabral HN, Bishop MJ (2011) Resistance among wild invertebrate populations to recurrent estuarine acidification. Estuar Coast Shelf Sci 93: 460–467.
- 58. Pietrock M, Marcogliese DJ (2003) Free-living endohelminth stages: at the mercy of environmental conditions. Trends Parasitol 19: 293–299.
- 59. Al Kurashi M, Eastick FA, Kuchipudi SV, Rauch C, Madouasse A, et al. (2011) Influence of culture medium pH on internalization, growth and phenotypic plasticity of Neospora caninum. Vet Parasitol 177: 267–274.
- 60. MacLeod CD, Poulin R (2012) Host-parasite interactions: a litmus test for ocean acidification? Trends Parasitol 28: 365–9.
- 61. Mackenzie CL, Ormondroyd GA, Curling SF, Ball RJ, Whiteley NM, et al. (2014) Ocean warming, more than acidification, reduces shell strength in a commercial shellfish species during food limitation. PLOS ONE 9(1): e86764.
- 62. Houghton JT, Ding Y, Griggs DJ, Nogouer M, van der Linden PJ, et al. (editors) (2001) Climate change 2001: the scientific basis. Contribution of Working Group I to the Third Assessment Report of the International Panel on Climate Change. Cambridge: Cambridge University Press. 881 p.
- 63. Lewis E, Wallace D (1998) Program developed for CO2 System calculations. Oakridge National Lab, US Department of Energy, Oakridge. 38 p.
- 64. Mehrbach C, Culberson CH, Hawley JE, Pytkowicz RM (1973) Measurements of the apparent dissociation constants of carbonic acid in seawater at atmospheric pressure. Limnol Oceanogr 18: 897–907.
- 65. Dickson AG, Millero FJ (1987) A comparison of the equilibrium constants for the dissociation of carbonic acid in seawater media. Deep Sea Res 34: 1733–1743.
- 66. Mortensen SH, Glette J (1996) Phagocytic activity of scallop (Pecten maximus) haemocytes maintained in vitro. Fish Shellfish Immunol 6: 111–121.
- 67. Pipe RK (1992) Generation of reactive oxygen metabolites by the haemocytes of the mussel Mytilus edulis. Dev Comp Immunol 16: 111–122.
- 68. Walsh PS, Metzger DA, Higuchi R (1991) Chelex 100 as a medium for simple extraction of DNA for PCR-based typing from forensic material. Biotechniques 10: 506–513.
- 69. Lynch SA, Mulcahy MF, Culloty SC (2008) Efficiency of diagnostic techniques for the parasite, Bonamia ostreae, in the flat oyster, Ostrea edulis. Aquaculture 281: 17–21.
- 70. Shi SR, Cote RJ, Wu L, Liu C, Datar R, et al. (2002) DNA extraction from archival formalin-fixed, paraffin-embedded tissue sections based on the antigen retrieval principle: heating under the influence of pH. J Histochem Cytochem 50(8): 1005–1011.
- 71. Inoue K, Waite JH, Matsuoka M, Odo S, Harayama S (1995) Interspecific variation in adhesive protein sequences of Mytilus edulis, M. galloprovincialis and M. trossulus. Biol Bull 189: 370–375.
- 72. Weisberg WG, Barns SM, Pelletier DA, Land DJ (1991) 16S ribosomal DNA amplification for phylogenetic study. J Bacteriol 173: 697–703.
- 73. Kawai M, Matsutera E, Kanda H, Yamaguchi N, Tani K, et al. (2002) 16S ribosomal DNA-based analysis of bacterial diversity in purified water used in pharmaceutical manufacturing processes by PCR and denaturing gradient gel electrophoresis. Appl Environ Microbiol 68: 699–704.
- 74. Zhang H, Bhattacharya D, Lin S (2005) Phylogeny of dinoflagellates based on mitochondrial cytochrome b and nuclear small subunit rDNA sequence comparisons. J Phycol 41: 411–420.
- 75. Le Roux F, Lorenzo G, Peyret P, Audemard C, Figueras A, et al. (2001) Molecular evidence for the existence of two species of Marteilia in Europe. J Eukaryot Microbiol 48: 449–454.
- 76. Renault T, Stokes NA, Chollet B, Cochennec N, Berthe F, et al. (2000) Haplosporidiosis in the Pacific oyster Crassostrea gigas from the French Atlantic coast. Dis Aquat Organ 42: 207–214.
- 77. Molloy DP, Giambérini L, Stokes NA, Burreson EM, Ovcharenko MA (2012) Haplosporidium raabei n. sp (Haplosporidia): a parasite of zebra mussels, Dressina polymorpha (Pallas, 1771). Parasitology 139: 463–477.
- 78. Shaw BL, Battle HI (1957) The gross and microscopic anatomy of the digestive tract of the oyster Crassostrea virginica (Gmelin). Can J Zoolog 35: 325–347.
- 79. Stevens A, Chalk BT (1996) Pigments and minerals. In: Bancroft JD, Stevens A, editors. Theory and practice of histological techniques, Fourth edition. London: Churchill Livingstone. pp. 243–267.
- 80. Mayrand E, St-Jean SD, Courtenay SC (2005) Haemocyte responses of blue mussels (Mytilus edulis L.) transferred from a contaminated site to a reference site: can the immune system recuperate? Aquac Res 36(10): 962–971.
- 81. Renwrantz L, Siegmund E, Woldmann M (2013) Variations in hemocyte counts in the mussel, Mytilus edulis: Similar reaction patterns occur in disappearance and return of molluscan hemocytes and vertebrate leukocytes. Comp Biochem Physiol A Mol Integr Physiol 164(4): 629–37.
- 82. Smith VJ, Chisholm JRS (1992) Non-cellular immunity in crustaceans. Fish Shellfish Immunol 2: 1–31.
- 83. Cheng W, Chen J-C (2000) Effects of pH, temperature and salinity on immune parameters of the freshwater prawn Macrobrachium rosenbergii. Fish Shellfish Immunol 10(4): 387–91.
- 84. Malham SK, Cotter E, O'Keeffe S, Lynch SA, Culloty SC, et al. (2009) Summer mortality of the Pacific oyster, Crassostrea gigas, in the Irish Sea: The influence of temperature and nutrients on health and survival. Aquaculture 287(1–2): 128–138.
- 85. Sokolova IM, Frederich M, Bagwe R, Lannig G, Sukhotin AA (2012) Energy homeostasis as an integrative tool for assessing limits of environmental stress tolerance in aquatic invertebrates. Mar Environ Res 79: 1–15.
- 86. Ellis RP (2013) The impact of ocean acidification, increased seawater temperature and a bacterial challenge on the immune response and physiology of the blue mussel, Mytilus edulis. PhD Thesis. University of Plymouth, UK.
- 87. Hooper C, Day R, Slocombe R, Handlinger J, Benkendorff K (2007) Stress and immune responses in abalone: limitations in current knowledge and investigative methods based on other models. Fish Shellfish Immunol 22: 363–379.
- 88. Fisher SW, Auffret M, Balouet G (1987) Response of European flat oyster (Ostrea edulis) hemocytes to acute salinity and temperature changes. Aquaculture 67: 179–190.
- 89. Lengeler J, Drews G, Schlegel H (2009) Biology of the prokaryotes. Google ebook ISBN: 978-1-4443-1330-7. Hoboken: John Wiley & Sons. 984 p.
- 90. Robledo JAF, Santarém MM, Figueras A (1994) Parasite loads of rafted blue mussels (Mytilus galloprovincialis) in Spain with special reference to the copepod, Mytilicola intestinalis. Aquaculture 127: 287–302.
- 91. Nappi AJ, Christensen BM (2005) Melanogenesis and associated cytoxic reactions: applications to insect innate immunity. Insect Biochem Molec 35: 443–59.
- 92. Katz ML, Robison WG Jr (2002) What is lipofuscin? Defining characteristics and differentiation from other autofluorescent lysosomal storage bodies. Arch Gerontol Geriatr 34: 169–84.
- 93. Jung T, Hohn A, Grune T (2010) Lipofuscin: detection and quantification by microscopic techniques. Methods Mol Biol 594: 173–193.
- 94. Carballal MJ, Villalba A, Lopez C (1998) Seasonal variation and effects of age, food availability, size, gonadal development, and parasitism on the hemogram of Mytilus galloprovincialis. J Invertebr Pathol 72: 304–312.
- 95. Paillard C, Allam B, Oubella R (2004) Effect of temperature on defence parameters in Manila clam Ruditapes philippinarum challenged with Vibrio tapetis. Dis Aquat Organ 59: 249–262.
- 96. Rohr JR, Schotthoefer AM, Raffel TR, Carrick HJ, Halstead N, et al. (2008) Agrochemicals increase trematode infections in a declining amphibian species. Nature 455: 1235–1250.
- 97. Palumbi S (2001) Humans as the world's greatest evolutionary force. Science 293: 1786–1790.
- 98. Lafferty KD (1997) The ecology of parasites in a salt marsh ecosystem. In: Beckage NE, Zuk M, editors. Parasites and Pathogens: Effects on Host Hormones and Behavior. New York: Chapman and Hall. pp. 316–332.
- 99. Hay SI (2002) Hot topic or hot air? Climate change and malaria resurgence in East African highlands. Trends Parasitol 18: 530–534.
- 100. Rogers DJ, Randolph SE (2006) Climate change and vector-borne diseases. Adv Parasitol 62: 345–381.
- 101. Rohr JR, Halstead NT, Raffel TR (2011) Modelling the future distribution of the amphibian chytrid fungus: the influence of climate and human-associated factors. J Appl Ecol 48: 174–176.
- 102. Food and Agriculture Organization (FAO) (2012) The State of World Fisheries and Aquaculture (SOFIA). FAO, Rome.
- 103. Shumway SE, Davis C, Downey R, Karney R, Kraeuter J, et al. (2003) Shellfish aquaculture - In praise of sustainable economies and environments. World Aquac 34: 15–19.