Physiological responses of the ghost shrimp Neotrypaea uncinata (Milne Edwards 1837) (Decapoda: Thalassinidea) to oxygen availability and recovery after severe environmental hypoxia

Comparative Biochemistry and Physiology, Part A 189 (2015) 30–37

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Physiological responses of the ghost shrimp Neotrypaea uncinata (Milne Edwards 1837) (Decapoda: Thalassinidea) to oxygen availability and recovery after severe environmental hypoxia Félix P. Leiva a,b, Mauricio A. Urbina c, Juan Pablo Cumillaf a, Paulina Gebauer b, Kurt Paschke a,⁎ a b c

Instituto de Acuicultura, Universidad Austral de Chile, P.O. Box 1327, Puerto Montt, Chile Centro de Investigación i~mar, Universidad de Los Lagos, Puerto Montt, Chile Department of Biosciences, College of Life and Environmental Sciences, University of Exeter, Exeter, United Kingdom

a r t i c l e

i n f o

Article history: Received 21 February 2015 Received in revised form 8 July 2015 Accepted 16 July 2015 Available online 26 July 2015 Keywords: Ghost shrimp Neotrypaea uncinata Ecophysiology Hypoxia tolerance

a b s t r a c t Hypoxia is a common and widespread phenomenon in aquatic ecosystems, imposing a significant challenge for the animals that inhabit such waters. In different habitats, however, the characteristics of these hypoxic events may differ, therefore imposing different challenges. We investigated the tolerance of adult ghost shrimp Neotrypaea uncinata (an intertidal mudflat dweller) to different partial pressures of oxygen (pO2), severe hypoxia (2 kPa) and recovery from hypoxia after different exposure times, mimicking the natural tidal cycle (6 h and 12 h). We calculated critical oxygen tension and categorize the adult ghost shrimps as oxyregulators (R value = 75.27%). All physiological measurements (metabolic rate, oxyhemocyanin, hemolymph protein and lactate concentrations) were affected by exposure to low partial pressures of oxygen, but most of them recovered (with exception of metabolic rate) control values (21 kPa) after 6 h under normoxic conditions. Low metabolic rate, high release of hemolymphatic proteins and anaerobic metabolism are suggested as response mechanisms to overcome hypoxic events during low tide. © 2015 Published by Elsevier Inc.

1. Introduction Owing to its presence and prevalence in aquatic and terrestrial habitats, hypoxia has long been a topic of interest. Hypoxia is defined as a shortage of oxygen (O2) that can be environmental or functional in origin (Farrell and Richards, 2009). Although there are terrestrial zones where partial pressure of oxygen (pO2) is greatly reduced such as the Himalayas, Alps and Andes, hypoxia in the aerial medium is generally uncommon (Schmidt-Nielsen, 1997). In aquatic environments, however, hypoxia is frequent and usually unpredictable. Although in some habitats such as the oxygen minimum zones (OMZs), hypoxic conditions are quite stable; in other habitats, hypoxia can also be unstable and predictable as dictated by the tidal cycle, i.e. intertidal mudflats (Childress and Seibel, 1998; Seibel, 2011). In coastal habitats and particularly in the marine intertidal zone, aerial and aquatic environments fluctuate on a periodic basis depending on the tide. Under a semidiurnal tidal cycle, intertidal zones are once or twice a day exposed to air or underwater. As a consequence, inhabitants of this zone must not only face dramatic changes in salinity, temperature, air

⁎ Corresponding author. E-mail addresses: [email protected] (F.P. Leiva), [email protected] (M.A. Urbina), [email protected] (J.P. Cumillaf), [email protected] (P. Gebauer), [email protected] (K. Paschke).

http://dx.doi.org/10.1016/j.cbpa.2015.07.008 1095-6433/© 2015 Published by Elsevier Inc.

exposure and desiccation but also must face drastic changes in pO2 (Willmer et al., 2000). Intertidal pools are the best example of such variations, where O2 saturation could range from 0% during the night to 400% during the day (Richards, 2011). Furthermore, it has also been reported that oxygen can only penetrate a few centimetres down into the sediment (Ziebis et al., 1996) exacerbating hypoxic conditions for the infauna. Notwithstanding these challenging conditions, there are some taxa able to live in such environments. Some marine invertebrates inhabiting intertidal zones are well-known for their ability to tolerate severe hypoxic or even anoxic events (Herreid, 1980; DeFur, 1988; McMahon, 1988; Grieshaber et al., 1994; Burnett and Stickle, 2001). Thalassinidean crustaceans are one of such groups inhabiting these environments. They live in burrows constructed in sandy or muddy sediments where hypoxic events are frequent and, consequently, they have developed a certain degree of hypoxia tolerance (for a review, Atkinson and Taylor, 2005). For example, pO2 as low as 0.65 kPa have been documented in burrows inhabited by Callianassa truncata (Ziebis et al., 1996). Such an ability would probably require adaptations at the behavioural, physiological and molecular levels. In fact, they can change their ventilatory behaviour while inside their burrows (Farley and Case, 1968; Miller et al., 1976; Torres et al., 1977; Astall et al., 1997), regulate metabolism over a wide range of pO2s (Thompson and Pritchard, 1969; Felder, 1979; Paterson and Thorne, 1995), have a high hemocyanin-O2 (Hc-O2) binding affinity (Miller et al., 1976; Miller and Van Holde, 1981; Taylor et al., 2000) and the ability to generate ATP by anaerobic metabolism to satisfy metabolic demands

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(Pritchard and Eddy, 1979; Zebe, 1982; Anderson et al., 1994; Holman and Hand, 2009). Neotrypaea uncinata, commonly known as the ghost shrimp, is a widely distributed thalassinidean. This species has been reported to live from the south of Mexico (~15°N) to the Peninsula de Taitao, Chile (47°S) (Thatje, 2003). In the south of Chile, N. uncinata, commonly known as “nape” by the fishermen (as it is used as a fish bait), is abundant in most sandy/muddy beaches. At this latitude, tides are typically semidiurnal with a period of approximately 12.4 h and maximum amplitude at spring tides of about 7 m. In addition, characterized by a low coastal bathymetric slope in certain locations, the intertidal zone of these beaches inhabited by N. uncinata can extend for several kilometres (i.e. Pelluhuin beach, Puerto Montt, Chile). In these natural settings, the tidal cycle plays a key role in modulating the periodicity of hypoxic events, presenting a natural experimental setup. With the exception of Astete-Espinoza et al. (2010), who evaluated the simultaneous effect of a bopyrid parasite and hypoxia on hemolymphatic parameters, no other study has explored the physiological responses that underline the ability of N. uncinata to cope with cyclic/tidal-related hypoxic events. The present study aimed to explore physiological responses that allow N. uncinata to thrive in an environment with such variable oxygen availability. Since both hypoxic events (low tide) and recovery (high tide) are controlled by the same driver and are time matched, we hypothesized that (i) N. uncinata exhibit a high regulatory capacity as environmental oxygen decrease and (ii) recovery from severe hypoxia should occur during a period determined by the high tide (i.e. 6 h). The physiological status and metabolism of N. uncinata exposed to different environmental pO2, and during different recovery times were measured. Critical partial pressure of oxygen (pcrit) and oxy-regulatory capacity were also assessed and used as an indicator of the hypoxia tolerance of this species. 2. Materials and methods 2.1. Animal collection and maintenance Adult intermoult male ghost shrimp were collected during low tide from the intertidal zone of Puntilla Tenglo, Puerto Montt, Chile (41°28′ S; 72°56′W). Shrimps were extracted from their burrows following the “Kiwi method” described by Torres et al. (1977). Briefly, the method consists of walking in circles (and gentle jumps), around the animal’s burrow. This action liquefies the muddy sand, causing animals to escape to the surface where they can be picked up by hand. The animals were placed in plastic containers with sand from the collection site, and transported to the Laboratory of Crustacean Ecophysiology (LECOFIC) at the Universidad Austral de Chile, Puerto Montt, Chile. In order to prevent aggression, the shrimps were kept in cylindrical PVC refuges (diameter 7.5 cm and 20 cm length) at a density of 25 shrimps per tank (35 L). Shrimps were maintained with constant flow of unfiltered seawater (32.3 ± 0.2 PSU), at 11.5 ± 0.2 °C, gentle aeration and 12 h:12 h light/ dark photoperiod inside a temperature-controlled room. Daily mortality was recorded (2% during the first three days in the laboratory, but no further mortality during either acclimation or experiments), and temperature and salinity were checked daily with a conductivity/temperature meter (WTW Cond 330i). Shrimps were fed daily with commercial krill flakes food (Tropical®). In order to avoid overlapping of oxygen consumption rate with a potential tide-related rhythm (Leiva et al. in preparation), shrimps were acclimated to these conditions for three weeks and feeding was stopped two days before the experiments began. Only intermoult males, with no missing appendages, and free from bopyrid parasites on the gills were used in the experiments. 2.2. Experimental design All experiments were conducted inside a temperature- and lightcontrolled room at the same conditions described above for animal maintenance, and using UV sterilized and filtered (1 μm) seawater.

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Series 1: Physiological responses to different pO2(s) and after recovery from severe hypoxia. A total of 42 shrimps, wet mass 15.72 ± 1.44 g (mean ± SD), CL 20.5 ± 0.7 mm, were randomly allocated to one of the five different experimental pO2(s): 21.11 (n = 6), 12.01 (n = 6), 7.65 (n = 6), 4.22 (n = 6) and 1.73 kPa (n = 18) (onwards referred as 21, 12, 8, 4 and 2 kPa, respectively). Shrimps were individually placed in 5 L hermetic respirometric chambers and exposed for 6 h to each respective experimental treatment, while oxygen consumption rate was quantified (see section Oxygen consumption rate (MO2) for more details). Experimental pO2(s) were previously prepared by bubbling N2 and leaving the water to equilibrate for 30 min prior to use. Chambers were submerged in a controlled-temperature bath at 11.5 ± 0.1 °C. Water oxygen content was measured at the beginning and at the end of the incubation period (6 h) for MO2 calculation (see section Oxygen consumption rate (MO2)). Thereafter, all shrimps allocated to the treatments 21, 12, 8, 4 kPa and a subset of only 6 shrimps of the ones allocated to 2 kPa were gently handled and a hemolymph sample was withdrawn for further analysis (see section Hemolymph sampling and analysis for details). Recovery was then evaluated in the remaining 12 shrimps that were exposed to 2 kPa. After the exposure to severe hypoxia, chambers were flushed with normoxic water (21 kPa). Subsequently, a subset of 6 shrimps was allowed to recover for 6 h in normoxia while the remaining (n = 6) shrimps were left to recover for 12 h in similar normoxic conditions. After their respective exposure and recovery periods, shrimps were sampled as previously described for the first group of shrimps. Oxygen water content was also quantified during recovery in order to determine MO2. In the treatment of 12 h of recovery, the chamber was flushed after 6 h and therefore water renewed, in order to avoid any potential accumulation of metabolic end products. Series 2: Effect of severe hypoxia and recovery time. The effect of severe hypoxia in ghost shrimp physiology was further evaluated by using a different set of 24 shrimps, wet mass 14.73 ± 1.40 g (mean ± SD), CL 20.0 ± 1.4 mm. Shrimps were randomly allocated to one of the four experimental treatments (each n = 6): normoxia 24 h (control, 21 kPa), severe hypoxia 12 h (2 kPa), severe hypoxia 12 h + recovery 6 h (21 kPa) and severe hypoxia 12 h + recovery 12 h. All experimental conditions, MO2 determinations and hemolymph sampling were conducted following the same procedures and under the same conditions as described above as for Series 1.

2.3. Oxygen consumption rate (MO2) Oxygen consumption rate was quantified by closed respirometry at each of the oxygen levels mentioned above (Series 1 and Series 2). Ghost shrimps were individually incubated in 5 L respirometric chambers with seawater previously UV sterilized and filtered (1 μm at 11.5 °C and 32 PSU). Animals were carefully introduced into the respirometric chambers, and subsequently left to acclimate to the respiratory chambers for at least 6 h prior the start of the experiments. During this period, all chambers were supplied with a constant flow (1 L h−1) of normoxic water coming from a reservoir. After this acclimation, the water in the chambers was replaced with the water at the corresponding oxygen level depending on treatment. This was achieved by flushing the chamber with water coming from a reservoir previously set at the corresponding oxygen level. Oxygen concentration was then measured before and after the incubation period (6 h) using an optic sensor connected to a Microx TX3 AOT (PreSens GmbH, Germany) oxygen meter previously calibrated in air (100% saturation) and with 5% sodium sulphite solution (0% saturation). Simultaneously, three chambers without shrimps, but treated identically, were also included during each MO2 determination to correct for potential bacterial oxygen consumption. MO2 were expressed as μmol of O2 per hour per gram dry weight (μmol O2 h−1 g−1). Animals were quickly rinsed with distilled water and dried for 48 h at 60 °C in a Memmert UFE 500 oven. Thereafter,

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the individual dry weight was determined using a semi-micro balance (Precisa GmbH, Precision 10 μg). 2.4. Calculation of pcrit The pcrit was calculated using all the individual MO2 values obtained at each experimental pO2 and adjusting a segmental linear regression. This allowed us to calculate the breakpoint (pcrit) when there was a sudden change in the response variable (Y: MO2) as a function of the independent variable (X: external pO2). An equation for each segment was calculated and pcrit was obtained using the intersection of the two linear regressions (Toms and Lesperance, 2003). 2.5. Hemolymph sampling and analysis Hemolymph samples (~ 500 μL) were withdrawn through the arthrodial membrane at the base of the fourth walking leg using a precooled 1 mL disposable syringe. Hemolymph was then quickly transferred to pre-cooled Eppendorf® microtubes (1.5 mL), and immediately frozen at −80 °C in an ultrafreezer (Thermo Scientific™, Forma Series 700) until analysis (Danford et al., 2002). Oxyhemocyanin concentration (Oxy-Hc) was calculated spectrophotometrically following previous protocols with slight modifications (Urbina et al., 2013). Briefly, 30 μL of crude hemolymph were diluted in 970 μL ultra-pure water (Barnstead™, Easypure®II), placed in a 10 mm path cuvette (1 mL) and the absorbance at 335 nm was read in a spectrophotometer (Thermo Scientific Multiskan™). Total hemolymph protein (Hem-Prot) was determined by the Lowry method using the DC Protein Assay Kit Bio Rad®, modified for a microplate reader. A calibration curve was constructed using bovine serum albumin (BSA) as standard. For L (+)-lactate determination (Hem-Lact), 100 μL of hemolymph was centrifuged at 6800g for 10 min at 4 °C, Hem-Lact was then determined in the supernatant via a colorimetric method using a Lactate Assay Kit (Spinreact, S.A./S.A.U., Spain), following the manufacturer’s instructions. The method was adapted to a microplate reader using 5 μL of hemolymph supernatant and 250 μL of reagent solution. Each sample was analyzed in duplicate, incubated to room temperature and its absorbance at 505 nm was then read on a microplate photometer (Thermo Scientific Multiskan®). 2.6. Oxy-regulatory capacity Regulatory capacity of N. uncinata was further evaluated by using the “R” regulation value, following the method proposed by Alexander and McMahon (2004). Briefly, each individual MO2 value obtained along the entire pO2 gradient tested was expressed as a percentage of the highest MO2 values recorded in all shrimps and treatments. Therefore, MO2 data then fell between 0% and 100%. External pO2 was also expressed as a percentage (0%–100%) and data was plotted and the best fit regression chosen. The area under that curve was then calculated by integrating the best fit equation from 0 to 100 as lower and upper limits, and expressing the results as a percentage of the total area. Thus, an animal displaying perfect oxygen regulation would have an R value of 100%, while an animal exhibiting oxygen conformity would have an R value of 50% (Alexander and McMahon, 2004). Animals that present “R” values between 50% and 100% are then able to oxy-regulate to a certain extent; the closer to 100%, the lower their pcrit and the closer to 50% the lower their ability of oxy-regulate (high pcrit). 2.7. Data analysis All data are presented as mean ± standard deviation. Differences in MO2, Oxy-Hc and Hem-Prot between treatments (pO2, hypoxia and recovery) were tested by a one-way ANOVA, followed by a Tukey post hoc test. Normal distribution and homogeneity of variances were previously checked with Kolmogorov–Smirnov and Levene’s tests, respectively. When the data did not adjust to parametric assumptions (Hem-Lact),

a Kruskal–Wallis test was used, followed by a Student–Newman– Keuls (SNK) test. For the calculation of the R index, the best fit regression curve was chosen on the basis of r2. Differences were considered significant with a P value b 0.05 (Zar, 2010). 3. Results No mortalities were recorded in any of the experimental treatments. During hypoxia exposures, shrimps were mostly quiescent with only occasional movements of the appendages, or changing position inside the chamber. 3.1. Series 1: Physiological responses at different pO2, and after recovery from severe hypoxia N. uncinata oxy-regulated from normoxia (21 kPa) down to a pcrit of 8.46 ± 1.32 kPa (Fig. 1A, vertical dashed line). Below this level, oxygen consumption of N. uncinata decreased as external pO2 did. Significant differences were found in the MO2 among pO2 treatments (ANOVA F6,35 = 30.981; P b 0.001). MO2 in normoxia (21 kPa) averaged 5.39 ± 0.40 μmol O2 h−1 g−1, and ghost shrimps were able to maintain this rate down to 8 kPa (4.96 ± 0.37 μmol O2 h− 1 g− 1; Tukey test, P N 0.05; Fig. 1A). Below 8 kPa, however, MO2 decreased to 3.56 ± 0.36 μmol O2 h−1 g−1 at 4 kPa and further down to 3.11 ± 0.36 μmol O2 h−1 g−1 at 2 kPa (both P values b 0.05). During the first 6 h of recovery in normoxia, after 6 h of hypoxia exposure, MO2 remained elevated, 7.05 ± 0.41 μmol O2 h−1 g−1 (Tukey test, P b 0.05; Fig. 1A). However, after 12 h of recovery, the MO2 had returned to pre-hypoxia exposure values, 4.95 ± 0.27 μmol O2 h−1 g−1 (Tukey test, P N 0.05; Fig. 1A). Oxy-Hc showed significant differences between treatments (ANOVA F6,35 = 12.433; P b 0.05). However, no differences in Oxy-Hc values were found in shrimps exposed to 21, 12, 8 and 4 kPa, showing an average of 1.03 ± 0.07 mmol L−1 in this pO2 range (Tukey test, P N 0.05; Fig. 1B). At 2 kPa, Oxy-Hc increased to 1.62 mmol L− 1, a value 57% higher than at higher pO2(s) (Tukey test, P b 0.05; Fig. 1B). During recovery, after either 6 h or 12 h in normoxic water, Oxy-Hc values returned to pre-hypoxic exposure values (Tukey test, P N 0.05; Fig. 1B). The pO2 treatments had a significant effect on Hem-Prot (ANOVA F6,35 = 9.695; P b 0.001). Hem-Prot remained unchanged from normoxia (21 kPa) down to 8 kPa, and the average value in this range was 56.59 ± 1.00 mg mL− 1 (Tukey test, P N 0.05; Fig. 1C). At 4 kPa, although not significantly different, a 12% increase in Hem-Prot was observed (P N 0.05; Fig. 1C). The highest values in Hem-Prot were observed in shrimp exposed to 2 kPa, with a mean of 80.32 ± 12.23 mg mL−1, significantly different from the Hem-Prot observed at 21, 12 and 8 kPa (all P values b 0.05; Fig. 1C). When shrimps were returned to normoxic conditions during either 6 h or 12 h, Hem-Prot returned to values observed in the 21–8 kPa pO2 range (Tukey test, P N 0.05; Fig. 1C). Under different pO2 treatments, the Hem-Lact concentration was significantly affected (Kruskal–Wallis H6 = 22.57; P b 0.001). HemLact concentration remained unchanged from normoxia (21 kPa) to a lower pO2 level of 8 kPa, with average values of 5.81 ± 0.99 mmol L− 1 (all P values N 0.05; Fig. 1D). At lower pO2(s), HemLact increased significantly, reaching mean values of 11.54 ± 4.94 mmol L− 1 and 15.8 ± 3.18 mmol L− 1, for 4 kPa and 2 kPa, respectively (SNK test, P b 0.05; Fig. 1D). When ghost shrimp recovered in normoxic conditions (21 kPa) during either 6 h or 12 h, Hem-Lact values returned to values similar to those reported between 21 and 8 kPa pO2 (SNK test, P N 0.05; Fig. 1D). 3.2. Series 2: Effect of the length of severe hypoxia and recovery time Significant differences between the studied recovery times were found (ANOVA F3,20 = 39.533; P b 0.001). Oxygen consumption in shrimps maintained in hypoxia for 12 h decreased by 49.69%

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Fig. 1. Physiological responses of N. uncinata exposed to different pO2(s) and after recovery from severe hypoxia. (A) Oxygen consumption (MO2, μmol O2 h−1 g−1) and pcrit (vertical dashed line), (B) oxyhemocyanin (Oxy-Hc, mmol L−1), (C) hemolymph protein (Hem-Prot, mg mL−1) and (D) L(+) lactate concentration (Hem-Lact, mmol L−1). Secondary graph at the right shows respective physiological responses following 6 h and 12 h of recovery, from 6 h of severe hypoxia (2 kPa treatment) in normoxic water (open circles). Data is presented as mean ± standard deviation (n = 6). Significant differences are indicated by different letters (Tukey or SNK test, P b 0.05).

(2.76 ± 0.16 μmol O2 h− 1 g− 1; Tukey test, P b 0.05; Fig. 2A), compared to a value of 5.56 ± 0.80 μmol O 2 h − 1 g − 1 in normoxia. Then, during the first 6 h of recovery in normoxic water, MO 2 increased to 8.19 ± 1.02 μmol O2 h− 1 g− 1, significantly higher than both control and hypoxic values (Tukey test, P b 0.05; Fig. 2A). However, after 12 h of recovery in normoxic water, MO2 had returned to control values (Tukey test, P N 0.05). Oxy-Hc varied significantly with the different treatments (ANOVA F3,20 = 52.417; P b 0.001). Oxy-Hc increased significantly by 81% during exposure to 12 h of hypoxia (P b 0.05; Fig. 2B) reaching values of 1.67 ± 0.17 mmol L−1 compared to an average value of 0.92 ± 0.06 mmol L−1 in the control treatment (Fig. 2B). After either 6 h or 12 h of recovery in

normoxic water, Oxy-Hc values were similar to the control treatment (Tukey test, P N 0.05; Fig. 2B). Recovery time after severe hypoxia caused a significant effect in Hem-Prot (ANOVA F3,20 = 27.634; P b 0.01). Compared to control values (47.89 ± 7.84 mg mL−1), Hem-Prot increased by 75% during 12 h of exposure to hypoxia (83.86 ± 8.62 mg mL−1; Tukey test, P b 0.05; Fig. 2C). After both 6 h and 12 h of recovery, Hem-Prot returned to values similar to that of the control treatment (P N 0.05; Fig. 2C). Significant differences in Hem-Lact between experimental conditions were found (Kruskal–Wallis H3 = 9.545; P b 0.05). Hem-Lact concentration in shrimps exposed to severe hypoxia for 12 h was 1.2-fold higher (14.21 ± 8.14 mmol L−1) than the Hem-Lact concentration

Fig. 2. Physiological responses of N. uncinata to prolonged severe hypoxia (2 kPa; 12 h) and different recovery times. (A) Oxygen consumption (MO2, μmol O2 h−1 g−1), (B) oxyhemocyanin (Oxy-Hc, mmol L−1), (C) hemolymph protein (Hem-Prot, mg mL−1) and (D) L(+) lactate concentration (Hem-Lact, mmol L−1) of N. uncinata exposed to either normoxia (Control), hypoxia (H12) or hypoxia followed by 6 h (H12 + R6) and 12 h (H12 + R12) of recovery in normoxic water. Data is presented as mean + standard deviation (n = 6). Significant differences are indicated by different letters (Tukey or SNK test, P b 0.05).

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observed in normoxic conditions (SNK test, P b 0.05; Fig. 2D). After both 6 h and 12 h of recovery, Hem-Lact concentrations reached similar values to that reported in the control treatment (21 kPa) (SNK test, P N 0.05; Fig. 2D). 3.3. Oxy-regulatory capacity The capacity for oxygen regulation was calculated by the integration of the area under the best fit non-linear equation curve (Michaelis– Menten model; r2, 0.806; P b 0.01) RðpO2 Þ ¼

94:410 ðpO2 Þ ð8:467 þ ðpO2 ÞÞ

In the range of pO2 tested, N. uncinata showed an R value of 75.27%, further indicating that adults of this species have the ability to oxyregulate (Fig. 3). 4. Discussion Our results show that adult N. uncinata are hypoxia tolerant, regulating their oxygen consumption down to ~8.46 kPa and then increasing their reliance on anaerobic metabolism below that oxygen threshold. Although the pcrit of N. uncinata was higher than that of other members of the thalassinids ghost shrimps (see below), no mortalities were recorded at any of the experimental treatments and a quick recovery was observed. 4.1. Metabolic rate and the effect of environmental pO2 When standardized data to wet mass is used, the intertidal burrowing ghost shrimp, N. uncinata, showed a metabolic rate of 1.31 ± 0.097 μmol O2 h−1 g−1, similar to values reported in closely related species such as Callianassa californiensis (1.2 ± 0.040 μmol O2 h−1 g−1) (Thompson and Pritchard, 1969). The maintenance of low metabolic rates is certainly advantageous for inhabiting extreme and fluctuating environments, and therefore it could be hypothesized that this maintenance is functionally adaptive in thalassinids, allowing them to survive hypoxic periods as a result of fluctuating intertidal environments (Thompson and Pritchard, 1969; Seibel, 2011). Further support for this idea comes from findings in other crustaceans (Whiteley and Taylor, 2015) and from the low metabolic rates found in two species of mudfish inhabiting mudflat environments that frequently dry out (Urbina et al., 2014a, 2014b). As any other taxa, decapod crustaceans vary greatly in their ability to maintain internal homeostasis when exposed to environmental stressors (Schulte, 2014). With regards to their respiratory responses to low oxygen levels, animals can be classified as oxy-regulators or

Fig. 3. Regulation capacity of N. uncinata. The relationship between MO2 and external oxygen saturation, both expressed as a percentage (%). Regulation value “R” is indicated by the area under the curve, according to Alexander and McMahon (2004).

oxy-conformers. While oxy-regulators are able to maintain their oxygen consumption down to sub-optimal oxygen levels, oxy-conformers are not, and their metabolic rates decrease as environmental pO2 does. The pO2 at which the animal stops oxy-regulating and starts oxyconforming is called the critical pressure of oxygen pcrit (Prosser, 1991). A pcrit of 8.46 ± 1.32 kPa was calculated for N. uncinata, showing a clear ability to oxy-regulate above this oxygen threshold. This clear oxy-regulating pattern was further confirmed by the calculated “R” regulation value (Alexander and McMahon, 2004), which classify this species as an oxy-regulator (R value of 75.27%). However, the pcrit found for N. uncinata is somewhat high compared with other members of the infraorder, such as C. californiensis, C. truncata and Trypaea australiensis (Table 1). Differences in pcrit values have been explained by a variety of factors both biotic (animal size, activity level, moulting cycle, oxyhemocyanin) and abiotic (temperature, salinity), which affect the ability of marine organisms to respond to environmental hypoxia (for a review, see Herreid, 1980). It could be noted from Table 1 that the shrimps used in the present study were bigger than those used in previous reports in other species. Although this might explain the differences in pcrit found based on differences in the surface/volume ratio emerging from different sizes (Dejours et al., 1970), further research is needed to test this hypothesis. Another explanation is, however, that pO2 levels in N. uncinata habitat do not decrease below their pcrit of 8.46 ± 1.32 kPa and therefore there is no driving force for extending oxy-regulation to a lower pO2. In agreement, Reardon and Chapman (2010) showed a decrease in pcrit in the cichlid Pseudocrenilabrus multicolor reared under low pO2 compared with fish reared under high pO2. Although in situ evaluations of the pO2 inside the burrows of N. uncinata are not available to date, pO2 values between 6.5 and 7.8 kPa have been reported in burrows of Upogebia stellata (at 12 °C, Astall et al., 1997). The pcrit also suggests that at above this level of hypoxia, animals are able to fuel their ATP demand by aerobic metabolism (Pörtner and Grieshaber, 1993; Hochachka and Lutz, 2001; Hochachka and Somero, 2002). This is also in agreement with our findings, as anaerobic metabolism (as measured by Hem-Lact) only increased below the pcrit of N. uncinata. 4.2. Coping with hypoxia Although crucial to maintaining ATP production below pcrit, anaerobic metabolism is not very energy efficient and therefore, attempts are made to maintain oxygen uptake and delivery to the tissues. Apart from the respiratory response showed and discussed above, where N. uncinata regulates and maintains its oxygen consumption down to ~8.46 kPa, oxygen delivery can also be enhanced by increasing the concentration and affinity of the respiratory pigment (Gorr et al., 2010). In fact, increases in hemocyanin-O2 (Hc-O2) binding affinity during hypoxia have been previously documented (Booth et al., 1982; Hagerman and Uglow, 1985; Hagerman and Baden, 1988; Baden et al., 1990; Spicer and Baden, 2000, 2001). In the present study, N. uncinata showed a significant increase in the hemolymphatic proteins and oxyhemocyanin below pcrit, reaching significantly higher values at 4 kPa and 2 kPa, respectively. An increase in the Oxy-Hc and Hem-Prot was also evident in the second series of exposure to hypoxia for 12 h. In agreement with our results, a recent study in the same species also found an increase in the Oxy-Hc after hypoxia exposure (Astete-Espinoza et al., 2010). The values of Oxy-Hc reported by those authors after hypoxic exposure were twice as high as the values obtained in the present study (3.72 ± 0.18 mmol L−1 vs 1.58 ± 0.09 mmol L−1, respectively). Astete-Espinoza et al. (2010), however, did not find an increase in Hem-Prot, reporting an average value of 75.18 ± 2.20 mg mL-1, similar to the Hem-Prot values found in the present study during hypoxic exposure. Together, these results strongly suggest that Oxy-Hc increases in order to enhance oxygen carrying capacity. An increase in the concentration of hemocyanin in response to hypoxia has been extensively documented in crustaceans such as Carcinus

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Table 1 Critical pressure of oxygen pcrit (kPa) among different marine decapod crustaceans. Species

pcrit

Temperature

Salinity

Wet mass

Reference

Upogebia pugettensis C. californiensis

6.5 2.2 1.3–2.6 4.8 8.46 7.8–10.4 7.9–10.4 5.2 2 5.2 6.6 7.4 4–9

10 14.2 10 22 11.5 15 10 10 10 15 10 10 12

33 n.a. 33 35 32 n.a. 32 31 n.a. n.a. n.a. n.a. 31

3.4–8.2 10–15 5.3–8.7 2–6 15.72 ± 1.44 60–80 250–450 35–130 500–1500 8.8 19–26 19–26 3.3 ± 1.1

Thompson and Pritchard (1969) Torres et al. (1977) Thompson and Pritchard (1969) Paterson and Thorne (1995) This study Taylor (1976) Bradford and Taylor (1982) Hagerman and Uglow (1985) Morris and Taylor (1985) Swain et al. (1987) Zainal et al. (1992) Zainal et al. (1992) Paschke et al. (2010)

T. australiensis N. uncinata C. maenas Cancer pagurus N. norvegicus Palaemon elegans Parastacoides tasmanicus Munida rugosa Munida sarsi L. santolla

Temperature (°C), salinity (PSU) and wet mass (grams) of experimental organisms. Not available data are indicated by n.a.

maenas (Taylor and Anstiss, 1999), Crangon crangon (Hagerman, 1986), Nephrops norvegicus (Hagerman and Uglow, 1985), Callinectes sapidus (DeFur et al., 1990), Macrobrachium rosenbergii (Chen and Kou, 1998) and Lithodes santolla (Urbina et al., 2013). This process is believed to occur quickly after the need for hemocyanin is detected. Although synthesis of new respiratory proteins is energetically expensive (Mente et al., 2003), the enhanced oxygen uptake and delivery expected should compensate the extra cost involved in its synthesis in the long term (Urbina et al., 2013). However, under an unexpected or short hypoxic event, it is conceptually counterintuitive to think that synthesis of new respiratory proteins will occur as a first response to low pO2. Higher vertebrates, such as fish, for example, store red blood cells in the spleen, and are readily released when they are needed (i.e. low pO2) (Randall and Perry, 1992). This implies that vertebrates do not incur in the extra cost (ATP and therefore O2) of synthesizing new hemoglobin when it is most needed; instead, they only release the previously synthesized and stored hemoglobin. A somewhat similar mechanism has been suggested for penaeid decapods, suggesting that hemocyanin is synthesized as a pre- protein composed by 661 amino acids. This preprotein is then stored in vesicles in the hepatopancreas until it is needed and converted to the mature protein of 648 amino acids (Sellos et al., 1997). Potential hemocyanin storage organs are also discussed in the literature, for example, small hemal sinuses are also considered responsible for the synthesis and storage of hemocyanin (Paulus and Laufer, 1987). This hypothesis is particularly attractive in the case of N. uncinata, which is exposed to cycling–short-term hypoxia (6 h to 12 h), where the energetic costs involved in up-regulation, translation, synthesis and release of a protein would be reduced and at the same time, a quicker response to this condition is facilitated. Further research is needed to test this hypothesis. As other respiratory pigments, hemocyanin in shrimps is the main protein making up the hemolymph. Since its concentration has been reported to account for between 60% and 90% of the total protein dissolved in the hemolymph (Rosas et al., 2004), variations in its concentrations should be seen in the total dissolved proteins in the hemolymph. Our data only partially support this idea, as hemolymph proteins (4 kPa) increased earlier than Oxy-Hc (2 kPa). This mismatch between Oxy-Hc and Hem-Prot in the hemolymph, suggest that although Oxy-Hc has been either released or synthesized at 4 kPa, it cannot readily bind to O2 until a lower pO2 has been reached (2 kPa). This strongly suggests that Hc-O2 binding is aided by a cofactor that reaches high enough levels at 2 kPa, facilitating Hc-O2 binding and transport to the tissues. Several cofactors such as lactate, urate, calcium and magnesium have been suggested to have a positive effect in Hc-O2 binding (Booth et al.1982; Morris and Taylor, 1985; Lallier and Truchot, 1989; Taylor and Whiteley, 1989; Morris, 1990; Danford et al., 2002). Lactate increased at the two lowest pO2(s) in our study, and therefore its rise might be related to the increase in Oxy-Hc levels found at 2 kPa. Whether or not another cofactor released during hypoxia such as alanine or succinate was playing

a role, remains to be confirmed. Results from the recovery experiments further support the hypothesis that a cofactor was aiding the Hc-O2 binding at the lowest pO2, but that Hc might also have been available at higher O2 concentrations (see following section). 4.3. Hypoxic recovery Recovery periods, after a hypoxic event wherein anaerobic pathways are activated, are crucial to restore energy reserves, osmotic, ionic and acid–base balance, oxygen levels, remove toxic end products, oxidize anaerobic end products and resume protein synthesis, which as a whole enables homeostasis to be restored (Herreid, 1980; Ellington, 1983, 2001; Maciel et al., 2008). As a result of the amplitude and frequency of the tidal cycle in the natural habitat of N. uncinata, both hypoxic exposure and recovery times occur in the same semidiurnal cycle. Lactate is the main end product of anaerobic metabolism in crustaceans (Livingstone, 1991) and its production and concentrations have been found to be particularly high in thalassinids (Holman and Hand, 2009). During recovery, lactate can be recycled back into pyruvate, then into glucose and subsequently into glycogen to restore energy reserves. In fact, muscular gluconeogenesis, and its diffusion into the extracellular medium has been reported in Chasmagnathus (Neohelice) granulata (Maciel et al., 2008). Therefore, although glucose and glycogen were not quantified in the present study, the mobilization and use of carbohydrates as fuel during the tidal cycle of hypoxia/reoxygenation is likely to occur in N. uncinata. Several studies in crustaceans have shown a direct relationship between hypoxia duration and the magnitude of the oxygen debt accumulated as consequence of the anaerobic end products generated (Grieshaber et al., 1994; Vismann and Hagerman, 1996; Zou et al., 1996). Our study shows that in ghost shrimps previously exposed to severe hypoxia for either 6 h or 12 h, full recovery took longer than 6 h in normoxia, but was achieved during the first 12 h of reoxygenation. Although no previous studies have evaluated the effect of severe hypoxia and recovery in thalassinids, studies in other decapods such as the crab Mennipe mercenaria found that during recovery from 12 h of hypoxia at b1 kPa, approximately 50% of the accumulated lactate had been removed during the first 6 h of recovery in normoxic conditions (Albert and Ellington, 1985), similar to our findings. Future experiments assessing oxygen consumption during recovery from hypoxia with higher time resolution would be useful in order to determine more precisely when full recovery occurs. In conclusion, N. uncinata presents a high regulatory capacity, in agreement with the first hypothesis. The mechanisms used by N. uncinata involve an increase in the concentrations of Hem-Prot and Oxy-Hc at low pO2 (2 kPa). However, a mismatch between these two variables suggests that there is a change in the hemocyanin affinity for O2, and lactate may be involved as a cofactor. Consistent with our second hypothesis, with the exception of MO2, all physiological variables evaluated returned to

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