Comparative Biochemistry and Physiology, Part A 161 (2012) 327–336

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Comparative Biochemistry and Physiology, Part A

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Effect of air exposure on lysosomal tissues of Mytilus edulis L. from natural intertidalwild beds and submerged culture ropes

M. Brenner a,b,c,⁎, K. Broeg a, C. Wilhelm d, C. Buchholz a, A. Koehler a,c

a Alfred Wegener Institute for Polar and Marine Research (AWI), Bremerhaven, Germanyb Institute for Marine Resources (IMARE), Bremerhaven, Germanyc Jacobs University Bremen, Bremen, Germanyd Furtwangen University, Villingen-Schwenningen, Germany

⁎ Corresponding author at: Alfred Wegener Institute(AWI), Bremerhaven, Germany. Tel.: +49 471 4831 103

E-mail address: Matthias.Brenner@awi.de (M. Brenn

1095-6433/$ – see front matter © 2011 Elsevier Inc. Alldoi:10.1016/j.cbpa.2011.12.001

a b s t r a c t

a r t i c l e i n f o

Article history:Received 7 June 2011Received in revised form 1 December 2011Accepted 1 December 2011Available online 8 December 2011

Keywords:AutophagyBlue musselHypoxiaLysosomal membrane stabilityStress response

Blue mussels collected from suspended culture ropes and from three natural intertidal wild beds from differ-ent areas of the German Bight were tested for their ability to cope with hypoxic conditions. During the exper-iment mussels were exposed to air from 0 to 72 h. Mussels from all sampling sites displayed high tolerance toaerial exposure with moderate levels of mortality after 12 to 48 h of exposure. Lysosomal membrane stability(LMS), a biomarker of general stress, changed notably between minimum values after 12 h and maximumvalues after 24 h of aerial exposure in intertidal mussels. In contrast, labilization times of mussels from thehanging culture increased continuously up to 48 h of exposure. Intertidal mussels from the island of Heligo-land exhibited significantly decreased membrane stability after 72 h of air exposure, correlating to highermortality rates. Intertidal mussels, although adapted to daily aerial exposure in their natural environment,showed a similar pattern of mortality and lower LMS values during the experiment than mussels from thesuspended culture site. The increase of LMS values of mussels under hypoxic conditions at the beginning ofthe experiment at all sites was tested for the influence of macro-autophagic processes using immune label-ling techniques. With this approach it could be demonstrated that high LMS values significantly correlatewith low autophagic activity. However, hypoxic conditions do not enhance autophagic processes duringthe early periods of aerial exposure. Only at the end of the experiment, high values for autophagy weremeasured in mussels from an intertidal site accompanied with high mortalities. The results indicate thatautophagic processes are not involved in the early adaptive processes that enable the mussel to cope withperiods of aerial exposure.

© 2011 Elsevier Inc. All rights reserved.

1. Introduction

Since the early 1970s when the first biomonitoring programmeswere initiated, blue mussels have been the internationally acceptedindicator species for the assessment of contamination levels andtheir harmful effects on marine and estuarine biota (Goldberg,1975; UNEP, 2007). In addition to having a wide geographic distribu-tion, blue mussels are abundant, stationary, and easy to sample.Further, mussels respond to natural abiotic changes, such as e.g. salin-ity, tidal or temperature variations (Bayne et al., 1985; Shakhmatovaet al., 1991; Zaldibar et al., 2004) and biotic stress caused by parasites,changing food availability or the presence of pathogens (Robledo etal., 1995; Sarà et al., 1998; Galimany et al., 2008). Thus, mussels areable to reflect integrative responses to various stress factors (e.g.Robinson and Langton, 1980; Labarta et al., 1997; Lòpez et al.,2001). As intertidal sessile organisms, mussels have to cope with

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harsh and rapidly changing physical conditions during tidal cycles,which include thermal stress (Helmuth and Hofmann, 2001), desicca-tion (Moore et al., 1979), anoxia (Hole et al., 1995), and reduced foodavailability during periods of emersion. In contrast, subtidal musselsand mussels cultured on submerged ropes inhabit a more stable envi-ronment (Hunt and Scheibling, 2001) although differences betweenhigh and low tide impact factors such as food availability whichcould affect their physiology. As a result, some intertidal bivalve spe-cies such as the ripped mussel Geukensia demissa (Dillwyn) exhibit ahigher capacity to adapt to environmental changes more rapidly thansubtidal organisms (Charles and Newell, 1997). According to Mooreet al. (2006a) animals living in a fluctuating environment are moretolerant towards pollutants since changes in salinities and food avail-ability as well as periodical hypoxia enhance the autophagicprocesses.

Macroautophagy, usually referred to as autophagy, is a majorpathway for bulk degradation of cytoplasmic constituents and organ-elles (Munafó and Colombo, 2001). In this process, portions of the -cytoplasm are sequestered into double membrane vesicles, theautophagosomes, and subsequently delivered to the lysosome for

Fig. 1. Map of the German Bight showing the sampling sites. Three intertidal sam-pling sites at Bordumer Sand (BS), Lyster Strand at the island of Sylt (SY), and thedune of the island of Heligoland (HL) and from one suspended hanging culture atthe Niedersachsenbrücke (nearshore) near Wilhelmshaven in the Jade (JD) Bay weresampled in April/May 2008.

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degradation and recycling (Klionsky and Emr, 2000; Kuma et al.,2004). The process is a highly conserved evolutionary mechanismfound in diverse groups of animals including nematodes, fliesand mammals (Cuervo, 2004; Moore et al., 2006a). Autophagy isup-regulated under stress-inducing situations including hypother-mia, hypoxia, environmental factors such as an increase in salinity(Moore et al., 2007) or physiological changes in the mussel; allowingcells to temporarily self-sustain in periods when nutrients are re-stricted (Bergamini et al., 2003). The energy required is producedby the recycling of proteins, lipids and cell organelles (Levine,2005). In addition to the role of autophagy as a survival strategy(Moore et al., 2006b) it is also discussed as an important responseto remove oxidatively damaged proteins and organelles (Bergaminiet al., 2003; Cuervo, 2004).

Various genes encoding for autophagy-related proteins are re-quired for the formation of autophagosomes. These genes were iden-tified by genetic screens in yeast, but in recent years related homologshave been found in many different organisms including mammals(Klionsky et al., 2003). One homolog of the autophagy-related pro-teins in yeast is the microtubule-associated protein light chain 3(LC3). This protein is essential for the formation of autophagosomes(Tanida et al., 2004). LC3 exists in a cytoplasmic form (LC3-I) andassociated with the autophagosome membrane (LC3-II). The amountof LC3-II correlates with the extent of autophagosome formation andis therefore used as an autophagosomal marker (Kabeya et al., 2000;Tanida et al., 2004; Wu et al., 2006).

A well-established tool to assess the health status of mussels fromdifferent sites is the test for lysosomal membrane stability (LMS). Thelysosomal responses in molluscan digestive cells constitute one of themost accepted biomarkers world-wide for assessing the health statusof the species and its environment (Marigómez et al., 1996; ICES,2006, 2007, 2008; UNEP, 2007). These tools, integrating the effectsof various classes of pollutants and indicating toxicant-induced cellpathologies, can provide evidence of the degree of stress or diseaseaffecting the organism (Allen and Moore, 2004). Lysosomes respondto a broad range of pollutants with a significant increase in size (e.g.Moore et al., 1978; Lowe et al., 1981; Moore, 1988; Cajaraville et al.,1989; Viarengo et al., 1992) and labilisation of their membranes(Moore et al., 1978; Viarengo et al., 1992; Koehler et al., 2002;Marigómez and Baybay-Villacorta, 2003). It has been shown thatthis response is correlated to site-specific contamination levels(Broeg et al., 1999, 2002; Kagley et al., 2003; Sturve et al., 2005;Schiedek et al., 2006). In addition to chemical pollutants, naturalstress factors such as temperature, salinity, availability and qualityof food, tidal cycle, and habitat conditions may also influence the sta-bility of lysosomal membranes (e.g. Marigómez et al., 1991; Tremblayand Pellerin-Massicotte, 1997; Abele et al., 1998; McVeigh et al.,2006). A decrease of the labilisation period and lysosomal enlarge-ment in mussels has also been shown to occur under combined hyp-oxic and hyperthermic conditions (Tremblay and Pellerin-Massicotte,1997). In addition to anthropogenic and natural stressors also meta-bolic processes such as reproductive status, infection or disease mayinfluence lysosomal properties (Bayne et al., 1985; Bignell et al.,2008; Broeg, 2010). Izagirre (2007) showed changes of lysosomalmembrane stability in mussels from the high intertidal, beginningwith aerial exposure, and discussed these results as a response todigestion processes. Since numerous factors may affect size andmembrane stability of lysosomes, appropriate experimental designsand a fundamental knowledge of basic responses are necessary tofacilitate interpretations of results with respect to the anthropogenicor natural causes of the observed effects.

Many bivalve species have developed strategies to cope with oxy-gen depletion in their natural environments. Intertidal organisms areexposed to air every tidal cycle during low tide, whereas animalsliving in subtidal areas may have to cope with periodic oxygendepletion in their habitat due to hydrographic (e.g. reduced water

exchange) or anthropogenic influences (e.g. increased oxygen de-manding degradation processes due to euthrophication) (Oeschger,1990). The species Arctica islandica, inhabiting the muddy bottomsof the Baltic Sea, for example, undergo self-induced anaerobiosis,since they dig burrows in oxygen depleted sediments. In experimen-tal trials A. islandica survived for 55 days in oxygen deficient seawater (Theede et al., 1969). Some intertidal acclimatized bivalvesclose their valves, maintaining a small gap through which oxygencan diffuse into the mantle cavity. Species like Mytilus californicusreach values up to 80% of the immersed rate of oxygen uptake by gap-ing under air exposure, whereas Mytilus edulis generally close thevalves during time of aerial exposure (Widdows et al., 1979).

Closing valves lead to oxygen deprivation in the tissues, followedby a profound depression of energy demand and a change from aero-bic to anaerobic metabolism (Eertman et al., 1993; Oeschger andStorey, 1993). Under anaerobic conditions, the fermentation of carbo-hydrates such as glycogen becomes the main source for energy inbivalves (de Zwaan andWijsman, 1976). This anaerobic fermentationprocess is less efficient than aerobic oxidation but may still exceed12% of aerobic ATP production under normal submerged conditions(Widdows et al., 1979). Using these strategies the blue mussel cansurvive in oxygen depleted water for up to 35 days (Theede et al.,1969) and under aerial exposure for up to 20 days (Babarro and deZwaan, 2008). Natural or anthropogenic stressors increase the meta-bolic rates of mussels and concomitantly the energy demand, thus,reducing aerial survival time compared to unstressed mussels (deZwaan and de Kock, 1988).

The aim of this experimental study was to obtain deeper insightsinto the influence of long-term aerial exposure on the lysosomalmembrane stability of the digestive gland cells ofM. edulis dependingon sampling site and habitat conditions (intertidal/submerged). Inaddition, the participation of autophagic processes as underlyingmechanisms of the acclimatisation to tidal related aerial exposurewas studied.

2. Materials and methods

Mussels were sampled at four locations along the coast of theGerman Bight (Fig. 1). Sampling was conducted between the 20thof April and the 5th of May 2007. Three areas were wild mussel

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beds: Bordumer Sand (BS, high intertidal, Position 53° 30′ 00′′N; 08°06′ 00′′ E), island of Heligoland (HL, intertidal, Position 54° 11′ 00′′ N;07° 54′ 09′′ E) and Lister Strand from the Island of Sylt (SY, low inter-tidal, Position 55° 01′ 10′′ N; 08° 26′ 50′′ E).

The intertidal areas were sampled during low tide. The fourthlocation was a test facility where mussels grew nearshore on sus-pended artificial substrates at the Niedersachsenbrücke, an approx.1.300 m long cargo bridge located in the Jade Bay (JD, Position 53°35′ 00 N; 08° 09′ 00 E). The artificial substrates were attached toten harnesses hanging from the bridge with 20 m between eachharness. Each harness consisted of a 20 mm polypropylene ropewith an iron plate (5 kg) at its distal end, weighing down the sub-strates into the water column even at strong current velocities. Theupper part of the harness was attached to a steel beam between thepillars of the bridge. The polypropylene (pp) rope was strengthenedwith two swirls and shackles to prevent entanglement. The lowerswirl was placed one meter above the mean high water (mHW)level to prevent corrosion and fouling. To insure that the cultivatedmussels were submerged throughout the experiment, samples ofartificial test collectors were fixed to the pp-rope from one to approx-imately three meters below mean low water (mLW) level using seawater- and uv-resistant plastic binders. The deployed substrate sam-ples were about 15 cm in length and fixed every 20 cm to the rope.

All intertidal sampling sites showed typical nearshore salinitiesbetween 28‰ and 31‰ throughout the year during high tide (unpub-lished data).

2.1. Sampling design

Approximately 80 mussels with shell lengths between 3 and 5 cmwere transported in a cool boxwithout water (dry) from each samplinglocation to the laboratory. Transport times for mussels from thesampling location to the lab varied between 2 h (BS and JD), 6 h HLand 12 h from the island of Sylt (SY). In the lab, all mussels from eachsampling site were stored separately in aerated 30 L sea water aquariafor adaptation (salinity 31‰) at natural water temperature (10 °C)illuminated for 12 h. Mussels were kept for 7 days in the aquaria andwere fed on the second and fifth day (5 mL, DT's Plankton Form). Oneday after feeding the aquaria were resupplied with fresh sea water

After 7 days, all mussels from one sampling site were taken out oftheir aquarium and kept dry under air exposure at a constant temper-ature of 10 °C. Ten mussels were analysed immediately (0 h). Shellparameters of each mussel were measured to the nearest 0.1 mmusing a Vernier calliper. Mussels were opened, drained and totalwet weight determined. The digestive gland was dissected, weighedand directly frozen in liquid nitrogen using cryo-vials. Subsequently,the soft body was removed and both shell and soft body wereweighed (±0.01 g) separately. The procedure was repeated with an-other ten mussels, kept under air exposure for 12 h, 24 h, 48 h and72 h. Mortality was calculated following the daily inspection prior topreparation. Mussels were considered dead and excluded from theexperiment when they were open and squeezing did not lead to im-mediate closing of shells. Mortality was calculated after subtractingthe ten mussels used for preparation and analysis after 0 h, 12 h,24 h, 48 h and 72 h, respectively.

dead animals= number of mussels survived−10ð Þ�100 ð1Þ

2.2. Sectioning and tissue preparation for histochemical analysis oflysosomal membrane stability

Tissue sections of a constant thickness of 10 μm were obtainedusing a cryotome (Microm, HM 500) with chamber temperature of−25 °C. Sections were stored for a maximum of 24 h at −20 °Cuntil processed for histochemistry. The lysosomal membrane stability

test was performed according to the cytochemical method describedby Moore et al. (2004). Serial cryostat sections were incubated at37 °C in a 0.1 M citrate buffer, pH 4.5, containing 3% NaCl to destabi-lize the membrane for increasing time intervals (2–50 min). Afterthis acid and temperature labilization, sections were incubatedfor 20 min at 37 °C in a medium containing the substrate NaphtholAS-BI N-acetyl β-D-glucosaminide (Sigma) dissolved in 2-methoxyethanol and low-viscosity polypeptide (Polypep, Sigma) dissolvedin 0.1 M citrate buffer, pH 4.5 with 3% NaCl. During incubation, thelysosomal acid hydrolase N-acetyl β-D-hexosaminidase catalyses therelease of the Naphthol AS-BI group. In the next step, the NaphtholAS-BI group undergoes a post-coupling reaction with the diazoniumsalt Fast Violet B (Sigma) dissolved in 0.1 M phosphate buffer(pH 7.4) leading to an insoluble bright violet reaction product.Following the colour reaction, samples were rinsed in running tapwater, fixed in Baker's Formalin and dried overnight in the dark. Sub-sequently slides were mounted in Kaiser's glycerine–gelatine.

2.3. Analysis of lysosomal membrane stability (LMS)

The maximum reaction product for N-acetyl β-D-hexosaminidase(peak) was determined by an automated measurement of numberand percentage of dark stained lysosomes in the digestive tubulesusing computer assisted image analysis (Zeiss, KS 300) combinedwith a light microscope (Zeiss, Axioskop) at 400-fold magnification.A long destabilization period indicates high lysosomal membraneintegrity and vice versa.

2.4. Sample sectioning and tissue preparation for autophagy assessment

The samples with frozen digestive glands used for the LMS assess-ment were also used for the assessment of autophagy. For this analy-sis a commercially available antibody (Anti-LC3, Sigma Aldrich) usedin the detection of LC3 proteins was chosen. The principle reactivityto blue mussel tissues and the working concentration for the immu-noblotting was tested using samples from the 0 h and 72 h exposuregroups. Thereby two sections per sample for the control and test of10 μmwere obtained using the above described cryotome with cham-ber temperature of −25 °C. Slides were stored at −80 °C until pro-cessed for immune labelling. Sections were prepared for incubationusing a PAP pen, providing a hydrophobic barrier around a tissuesample on the slide to keep incubation reagents localized on the tis-sue sections. Incubation of sections was conducted at room tempera-ture in a dark and humid chamber by treating the samples three timesfor 5 min using 0.1 M PBS buffer (pH 7.2), subsequently three timesfor 10 min using 0.1% glycin-PBS buffer, followed by a 20 min incuba-tion using a 1% BSA-PBS solution. After this preparation the first anti-body (Anti-LC3, Sigma Aldrich) was applied (2 μg/mL 1% BSA-PBSsolution) for 4 h only on the test tissues. After the incubation slideswere rinsed five times for 5 min in a 0.1% BSA-PBS buffer. In a nextstep, a second fluorescing antibody (Alex Fluor 350, Invitrogen,Darmstadt, Germany) was applied. The stock solution was diluted(1:300) in a 1% BSA-PBS buffer. Samples were incubated for 2 h in adark and humid chamber using 200 μL of the antibody solution foreach test and control tissue. Slides were then rinsed 5 times for5 min in a PBS buffer, incubated for 15 min in 1% glutaraldehyde-PBS-solution before being rinsed again 3 times for 5 min in distilledwater. In a final step slides were dried and mounted using a non-fluorescent medium (Dako, Carpinteria, USA).

Slides containing control tissues for each site and the two treat-ment groups (0 h and 72 h) were treated identically, except that sam-ples were not incubated with the first antibody. After preparation allslides were stored cool (4 °C) and dark until analysed under a lightmicroscope.

The assessment of the test treatment revealed best fluorescingresults using a concentration of 2 μg/mL of the primary antibody.

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Since the trials showed a clear pattern of anti-LC3 reactivity to themussel tissues (no fluorescence in the controls and different intensi-ties of fluorescence in the test tissues), all samples (according to

site and treatment) were prepared thereafter following the processesdescribed above.

2.5. Autophagy analysis

Incubated slides were analysed with a light microscope (Zeiss,Axioskop) under 1000 fold magnification using non-fluorescentmmersion oil. Photos of the blue fluorescent areas indicating the pres-ence of autophagic proteins were taken, supported by the Axiovisonsoftware (Release 4.7, Carl Zeiss Vision, Munich, Germany). Evalua-tion of photos was conducted semi-quantitatively by ranging theoccurrence of fluorescence, resulting from the autophagy indicatingproteins, into 4 categories: 0 = no fluorescence, 1 = fluorescence,2 = moderate fluorescence and 3 = high fluorescence (Fig. 2).

2.6. Statistical analysis of data

All statistical analyses were conducted using the SigmaPlot 11.0and Statistica 9.0 software. The non-parametric Kruskal–WallisANOVA (SigmaPlot) was used to compare time of air exposure (0 h,12 h, 24 h, 48 h and 72 h) for peak 1 and 2 at all sites. This test wasfollowed by an all pair-wise multiple comparison procedure (Dunn'sMethod) to describe differences between the sites for peaks 1 and 2.In addition initially dissected mussels (0 h) were compared forpeak 1 and 2 specific to the four different sites. This procedure wasrepeated at the end of the experiment (72 h). Box–Whisker plotswere calculated for all sites and for both peaks and correlations be-tween the LMS and autophagic activities were conducted using theStatistica 9.0 software. Prior to the correlation all LMS values ofpeak 1 and 2 were assigned to four categories according to Viarengoet al. (2007). Categories for peak one are: 4 (b5 min labilisationtime)=irreversibly damaged; 3 (5 tob10 min labilisation time)=severely stressed; 2 (10 tob15 min labilisation time)=good condi-tion and 1 (≥15 min labilisation time)=healthy. Analogue categoriesfor peak 2 were set as: 4 (10 and 15 min labilisation time)=irrevers-ibly damaged; 3 (20 and 25 min labilisation time)=severelystressed; 2 (30 and 35 min labilisation time)=good condition and 1(>35 min labilisation time)=healthy.

3. Results

3.1. Mortality

At the beginning of the experiment n=66 (JD), n=85 (BS),n=80 (SY) and n=80 (HL) mussels were exposed to air. Under24 h of exposure the mortality was relatively low at all sites andvaried between 3% (BS) and 11% (SY) (Fig. 3). After 48 h of exposure,mortalities increased and varied between 5% (BS) and 24% (HL). Atthe end of the experiment after 72 h, more than 50% of the musselsfrom HL were dead, whereas the mortality at BS was only 20%. JDand SY showed intermediate mortalities of 38% and 32% after 72 h(Fig. 3).

3.2. Lysosomal membrane stability (LMS)

At the beginning of the experiment (0 h, Fig. 4a/b) LMS values forpeaks 1 and 2 varied between the sites. Mussels from the intertidalareas SY and BS showed the highest membrane stability, as indicatedby long destabilisation periods for peaks 1 and 2, whereas the mem-brane stabilities of mussels from the cultivation area JD and the

Fig. 2. a–d: Exemplary photos (400 fold magnification) of the 4 different (blue) fluo-rescing categories (a) category 0 = no fluorescence, (b) category 1 = fluorescence,(c) category 2 = moderate fluorescence and (d) category 3 = high fluorescenceresulting from the semi-quantitative analysis of autophagic processes after aerial ex-posure in the micro-tubuli of mussel's digestive gland.

Fig. 3. Mortality in % of mussels from 4 different sampling sites JD (white rhombus,n=66), BS (black squares, n=85), SY (black circles, n=80) and HL (black triangles,n=80) during the time of aerial exposure (0 h-72 h).

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intertidal area HL were lower. Fig. 5a–h shows the results for all airexposure times (0 h, 12 h, 24 h, 48 h and 72 h) of each sampling site(peak 1 and peak 2). At all intertidal areas (BS, SY and HL) the values

Fig. 4. a/b: Box–Whisker plots comparing the labilisation time [min] of peak 1 (a) andpeak 2 (b) of the lysosomal membrane stability test for all sampling sites (JD, BS, SYand HL) at the beginning of the experiment (t 0 h).

for peak 1 and 2 were highest after 24 h of air exposure (Fig. 5c–h). AtJD the highest labilization values were achieved even later after 48 hof air exposure (peak 1 and 2) (Fig. 5a/b). Significant differences weredetected at HL for peak 2 (Fig. 5h) between HL 00 vs. HL 24 (Pb0.05),HL 12 vs. HL 48 (Pb0.05) and HL 48 vs. HL 72 (Pb0.01). In Fig. 6a/bthe LMS values for all sites at the end (72 h) of the experiment aredisplayed. At intertidal sites labilisation values were lowest at theend of the experiment. A significant difference was detected forpeak 2 between JD vs. BS (Pb0.05).

3.3. Autophagic activity

The results of the immune labelling of LC3 proteins characteristicfor autophagic processes are shown in Fig. 7a–d. Mussels from JD(Fig. 7a) show higher autophagy compared to the mussels from theintertidal sampling sites (Fig. 7b–d). At all sites, autophagic processesremain unaffected by hypoxic conditions for up to 48 h of aerial expo-sure. Only by the end of the experiment did the autophagic activityincrease significantly at the intertidal site BS (P>0.05) (Fig. 7b+d).

3.4. Correlation of LMS and autophagic activity

The LMS values (assigned to four categories, see Section 2.6) andautophagic activity showed a significant negative correlation(n=34, Pb0.05) of peak 1 and 2 in the control group (0 h) at the be-ginning of the experiment, indicating that high LMS values correlatewith reduced autophagic activity. However, LMS values from allgroups regarding aerial exposure (12 h, 24 h, 48 h and 72 h) did notcorrelate with autophagic activity.

4. Discussion

Mussels are equipped with specialized mechanisms to cope withreduced oxygen availability under conditions such as exposure toair in intertidal areas or oxygen depletion in the water column.Exposed to air, some species are able to reduce the oxygen deficiencyby gaping, but typically bivalves reduce their energy demand andswitch from aerobic processes to anaerobic metabolism to maintaina minimum of energy supply (de Zwaan and Wijsman, 1976;Eertman et al., 1993). Reducing their demand for energy duringanaerobiosis is achieved by: halting food consumption and processing(de Zwaan and Eertman, 1996), by reducing heart-beat frequency,ventilation (Bayne, 1971) and energy consuming processes such asprotein-synthesis (Boutilier, 2001) and by the suppression of nucleartranscription (Larade and Storey, 2002). Further, anoxia tolerantmolluscs have large tissue stores of fermentable fuels, such as glyco-gen and selected amino acids (Larade and Storey, 2002) and producevolatile or less acidic end-products during fermentation that do notperturb cellular homeostasis (Isani et al., 1995). With these metabolicadaptations, some species e.g. Artica islandica are able to survive forweeks under anoxic conditions (Oeschger and Storey, 1993). Bluemussels, whose natural habitat includes sub- and intertidal areas,emerge periodically during low tide and are able to survive for daysunder conditions of air exposure or even weeks in oxygen depletedwaters (Theede et al., 1969).

This study covered a large geographic area encompassing differenttypes and sizes of water bodies and habitats, thereby including abroad spectrum of mussels varying in health depending on the differ-ent exposures to contaminants and the different acclimatisationlevels to tide-related aerial exposure. According to the database(MUDAB) of the Federal Maritime Hydrographic Agency (BSH,2009), the coastal and nearshore sites (BS, SY and JD) showed compa-rably high contamination loads in the water column and suspendedparticles, whereas concentrations around the island of Heligolandfurther offshore were lower.

Fig. 5. a–h: Box–Whisker plots comparing the labilisation time [min] of peak 1 and peak 2 of the lysosomal membrane stability test for all sampling sites (a) JD peak 1, (b) JD peak2, (c) BS peak 1, (d) BS peak 2, (e) SY peak 1, (f) SY peak 2, (g) HL peak 1 and (h) HL peak 2 in relation to tested times of hypoxia (0 h, 12 h, 24 h, 48 h, and 72 h). Differences(Kruskal–Wallis-Test/Dunn's Method) for peak 2 between HL 00 vs. HL 24 (Pb0.05), HL 12 vs. HL 24 (Pb0.05), and HL 48 vs. HL 72 (Pb0.01), are significant (*).

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Fig. 6. a/b: Box–Whisker plots comparing the labilisation time [min] of peak 1 (a) andpeak 2 (b) of the lysosomal membrane stability test for all sampling sites at the end ofthe experiment (t 72 h). Differences (Kruskal–Wallis-Test/Dunn's Method) for peak 2(b) between JD and BS are significant (P>0.05) (*).

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Although mussels from all sampling sites were acclimated for7 days in aquaria with clean sea water prior to the experiment,lysosomal membrane stability differed between the sites. Membranesof mussels taken from the intertidal areas were more stable at thebeginning of the experiment than those taken from the submergedcultivation site. At the end of the experiment, this pattern was re-versed and mussels from the submerged cultivation site showedhigher membrane stabilities.

During the experiment, labilization periods decreased at all inter-tidal sites between 0 and 12 h of aerial exposure followed by an in-crease of the labilization periods between 12 and 24 h. Changes oflysosomal membrane stabilities shortly after emersion were alsoreported by Izagirre (2007). The author concluded that digestion pro-cesses i.e. intracellular digestion, triggered by the tide, might be re-sponsible for the decrease in lysosomal membrane stability duringthe time of air exposure. The increase of membrane stability at all in-tertidal sites between 12 and 24 h of air exposure is followed by a de-cline of values until the end of the experiment. This patterncorresponds with the results of Moore et al. (2007), where a markeddecrease in lysosomal membrane stability was detected after 24 h ofair exposure. In contrast to the results from the intertidal sites, mus-sels from the cultivation plot showed a different pattern during thetime of aerial exposure. Here, a constant increase of membrane stabil-ity is displayed until 48 h of exposure, before membranes destabilizedat the end of the experiment. Interestingly, mussels from all sitesreached the maximum of membrane stability during the period ofaerial exposure.

The study of Moore et al. (2007) suggested that a stressful fluctu-ating environment triggers autophagic events leading to the

elimination of altered proteins or damaged cellular constituents,and the formation as well as accumulation of peroxidation end prod-ucts such as lipofuscin is minimised. This could explain the higher ly-sosomal membrane stability values measured at the beginning of theexperiment found in the intertidal mussels. However, these high LMSvalues were negatively correlated with autophagy, indicating lowerautophagic activities in the intertidal mussels compared to the sus-pended culture mussels. Autophagy was not the cause for changesin lysosomal membrane stability measured throughout the experi-ment as the autophagic activity continued to be low and did not cor-relate with LMS values during aerial exposure. These results mayindicate that enhanced autophagy does not directly account forhigher LMS values or, more likely, autophagy accounts for higher sta-bility, but these processes occur over time and could not be demon-strated using the approach presented in this study.

However prolonged periods of hypoxic conditions do not neces-sarily indicate additional stress, as mussels are able to recover e.g.their lysosomal integrity under anaerobic conditions. Yet clearlybeyond a certain length of time under conditions of aerial exposure(in this study after 48 h), the impact of anoxic conditions induce anotable increase in mortality. This enhanced mortality correspondedwith a decrease in the LMS at all three intertidal sites, where thelowest labilization values of the entire experiment were detected.Supporting these findings, the increased autophagic activity wasdetected at the end of the experiment at the intertidal site BS. Howev-er, since blue mussels can survive even longer than 72 h underhypoxic conditions (Theede et al., 1969; Babarro and de Zwaan,2008) a correlation between the decrease in LMS, autophagy andmortality at all other sites might occur even later.

In conclusion, “hypoxia” is not increasing cellular impairmentsleading to enhanced autophagic repair processes. Only in case ofsevere O2 limitations increase in autophagic activity and/or enhancedapoptosis occur. As described in several other studies autophagy is acellular response to starvation in many different organisms (e.g.Levine and Klionsky, 2004; Levine, 2005; Moore et al., 2006c; Toddeet al., 2009; Klionsky, 2010; etc.). However, mussels from theGerman Bight show, probably due to an enhanced eutrophicationparticularly in the coastal zones, high concentrations of glycogen incells of the digestive gland (Brenner, 2010). These high energyreserves may help to prevent starvation in the mussels even underprolonged emersion times as shown in this experiment. Togetherwith the ability of mussels to reduce metabolic rates and to switchfrom aerobic processes to anaerobic metabolism, mussels may beable to generate sufficient energy to sustain an adequate metabolicrate under long lasting hypoxic conditions.

Reducing the metabolic rate combined with fermentative energyproduction are probably the most important factors for mussel sur-vival under anoxic conditions, as has been described in other studies(e.g. Zurburg and Kluytmans, 1980; Hermes-Lima and Zenteno-Savín,2002; Babarro and de Zwaan, 2008; Anestis et al., 2010). Energy rec-lamation by autophagic recycling is obviously not a strategy used bymussels to adapt to hypoxia or anoxia, since protein-turnover is anATP consuming aerobic pathway (Boutilier, 2001) to which speciesare very sensitive during anaerobiosis (de Zwaan and Eertman, 1996).

This study shows that acclimatisation to aerial exposure by the in-tertidal mussels does not offer a relevant advantage under prolongedexposure conditions, as mussels from suspended cultures showedeven higher labilisation values compared to intertidal mussels after48 and 72 h. These findings indicate that the ability to cope withlong term anoxia is genetically determined rather than being a ques-tion of phenotypic acclimatisation. Interestingly, all mussels regard-less of origin and habitat properties showed low membranelabilisation values of 3 to 8 min for peak one, which did not differ sig-nificantly between sites. According to Viarengo et al. (2007) these an-imals would be considered as severely stressed, already exhibitingpathologies. Following this general classification our results indicate

Fig. 7. a–d: Box–Whisker plots showing autophagic activity at all sampling sites (a) JD, (b) BS, (c) SY and (d) HL according to the different times of areal exposure (0 h, 12 h, 24 h,48 h, and 72 h). Autophagic activity is displayed in four categories of fluorescence (0=no fluorescence, 1=fluorescence, 2=moderate fluorescence and 3=high fluorescence),resulting from the immune labelling of autophagy indicating proteins. Differences (Kruskal-Wallis-Test/ Dunn's Method) in the autophagic activity at BS (c) between BS 72 vs.BS 48, BS 72 vs. BS 24 and BS 72 vs. BS 12 were significant (P>0.05) (*).

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that the quality of the coastal waters of the German Bight—includingthe area around the island of Heligoland—poses a risk to the health ofits populations of blue mussels. Low labilization times were alsofound in mussels from the same sampling site (JD) over the wholeyear of 2007 and in mussels from intertidal wild beds (including BSand SY) of the German Bight in the year 2007 (Brenner, 2010).

In addition to the pollutant levels, the extent of parasite infesta-tions also varies between sites along the coast of the German Bight.The waters near the island of Heligoland have a low level of pollutants(BSH, 2009), but the mussels suffer, due to the existence of all para-sitic hosts needed to fulfil the life cycles of the different parasiticspecies in confined spaces, from mass infections (Voss, 2006). Thetwo remaining intertidal areas in this study were analysed for para-sites in 2007 (Brenner et al., 2009). As also shown in previous inves-tigations (Buck et al., 2005; Thieltges et al., 2006) these areasdisplayed high infestation rates by a large variety of parasitic species.Mussels from the hanging cultivated area in the Jade Bay had low in-festation rates and were infested by only one species (Brenner et al.,2009). Parasites function as a trigger of the mussels’ immune system.According to Veldhuizen-Tsoerkan et al. (1991) and Eertman et al.(1993) survival in air is reduced under additional stressful environ-mental conditions, as mussels which are subject to toxic substancesor natural stress have an increased metabolic rate (de Zwaan and deKock, 1988), leading to higher mortalities under aerial exposure (stresson stress response) (Smaal et al., 1991; Viarengo et al., 1995). Althougha correlation between parasite infestation rates and mortality could notbe proven using the approach in this study, the results follow the patternsof previous investigations. Mussels from the island of Heligoland, whichshowed the highest infestation rates, also showed the highest mortalityunder conditions of aerial exposure.

5. Conclusions and outlook

The results of this study show that intertidal organisms, such asblue mussels, have developed robust strategies to cope with anoxiawithout suffering severe metabolic stress. However, it remained diffi-cult to differentiate the relevance of the complex metabolic processesinvolved in phases of hypoxia delineated using this experimentalapproach. However, mussels are able to maintain the integrity of ly-sosomal membranes even under long term anoxic conditions, sincea sudden release of high amounts of hydrolytic enzymes due to mem-brane leakage, would result in self-digestion and death of the organ-isms. Thus, the key function of lysosomal membrane stability for theviability of the organism is highlighted. Furthermore, it could be dem-onstrated that mussels are even able to recover under anaerobic con-ditions using, most probably, fermentation processes for energyproduction. These energy generating processes are activated onlyunder prolonged hypoxic conditions—a twelve hour period is almosttwice the time of a tidal cycle in the German Bight (approx. 6 h and20 min). Cultivated mussels, which were never exposed to air previ-ously, are able to switch immediately to these likewise genetically de-fined alternative pathways.

Acknowledgments

The authors are grateful to the Niedersachsen Port Authorities(NPorts) who generously allowed us to use the cargo bridge at theJade Bay for deploying the mussel collectors. This work was realizedwithin the project FV 168 MytiFit financed by the Alfred Wegener In-stitute for Polar and Marine Research, Bremerhaven, Germany and

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the Senate of Construction, Environment, Traffic and Europe of theFederal State of Bremen, Bremen, Germany.

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