Although, Linares et al. (2015) did not evaluate the haemolymph AA of O. mimus, considering that glycogen was also accumulated in its muscle after feeding, we conjecture that as observed in O. maya, the muscle glycogen in O. mimus was synthesized from AA, reaching its maximum value 400 min after feeding (**Figure 2B**).

At the end of the digestive process (480 min after feeding), a DG pH reduction was reported in O. maya and O. mimus (Martínez et al., 2011), suggesting a release of digestive enzymes in preparation for a new digestive cycle. Those enzymes could be sent to the digestive tract, where a reduction of pH was also registered (Linares et al., 2015). Reduction of enzymatic activity could also indicate that enzymes in the new gastric juice were zymogens that require the chyme to be totally activated.

Following the histological dynamics of DG in O. maya, Martínez et al. (2011) also observed an increment of residual body density 360 min after feeding, indicating that the feces and cellular debris removal process reached its maximum level at that time. Posteriorly, all the activity in the digestive system was reduced, with low production of residual bodies in the DG cells indicating that digestive cycle had ended (**Figure 2A**). At that time, nutrient reserves were accumulated in wait for the next meal (Martínez et al., 2011; **Figure 2B**).

As was previously stated for O. vulgaris by Boucaud-Camou and Boucher-Rodoni (1983), is evident the digestive physiology of O. maya and O. mimus is a fast and strongly dynamic process. In adults, this process takes around 480 min to be completed, indicating that this type of animal should be fed at least every 8 h to maintain its health in captivity (Linares et al., 2015). At a semi-pilot scale, this feed protocol has been followed for more than 5 years (Rosas et al., 2014); adults of O. maya were fed every 8 h using fresh scraps of marine fish or fresh crab (Caamal-Monsreal et al., 2015) or a diet formulated to stimulate spawning in laboratory conditions (Tercero-Iglesias et al., 2015). Under these conditions the number of eggs spawned was quite similar to those observed in wild spawns (Vidal et al., 2014), indicating that laboratory animals fed every 8 h reach a similar healthy condition to those on the continental shelf of the Yucatán Peninsula, where this species lives (Avila-Poveda et al., 2016; Angeles-Gonzalez et al., 2017). O. maya and O. mimus are well adapted, as are the majority of cephalopod species, to digest a high-quality animal protein diet using a mix of acidic and alkaline enzymes. This allows them to efficiently obtain the energy and molecules necessary to maintain their physiological functions according to the environment where they live, as shown for the tropical (22–30◦C; O. maya) and temperate (14–22◦C; O. mimus) species.

#### AUTHOR CONTRIBUTIONS

CR, PG, AO, RM, and CC designed and ran the experiments in Mexico and Chile. CR, PG, PD, MM, CP, and AS, performed the laboratory analysis and processed the data. All authors contributed to write the paper.

# FUNDING

This research was partially financed by the project PAPIIT IT201117 from the DGAPA-UNAM to PG and the project PAPIIT IN219116 from DGAPA-UNAM to CR. Also thanks are given to Dirección General de Cooperación e Internacionalización of UNAM for support gave to TEMPOXMAR research net.

#### ACKNOWLEDGMENTS

We acknowledge the support from COST action project: COST Action FA1301.

# REFERENCES

digestibility of myofibrillar proteins. J. Agric. Food Chem. 56, 1488–1494. doi: 10.1021/jf072999g

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2017 Gallardo, Olivares, Martínez-Yáñez, Caamal-Monsreal, Domingues, Mascaró, Sánchez, Pascual and Rosas. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# The Role of the Cephalopod Digestive Gland in the Storage and Detoxification of Marine Pollutants

Ana P. Rodrigo and Pedro M. Costa\*

Environmental Toxicology Lab, MARE - Marine and Environmental Sciences Centre, Departamento de Ciências e Engenharia do Ambiente, Faculdade de Ciências e Tecnologia da Universidade Nova de Lisboa, Caparica, Portugal

The relevance of cephalopods for fisheries and even aquaculture, is raising concerns on the relationship between these molluscs and environmental stressors, from climate change to pollution. However, how these organisms cope with environmental toxicants is far less understood than for other molluscs, especially bivalves, which are frontline models in aquatic toxicology. Although, sharing the same basic body plan, cephalopods hold distinct adaptations, often unique, as they are active predators with high growth and metabolic rates. Most studies on the digestive gland, the analog to the vertebrate liver, focused on metal bioaccumulation and its relation to environmental concentrations, with indication for the involvement of special cellular structures (like spherulae) and proteins. Although the functioning of phase I and II enzymes of detoxification in molluscs is controversial, there is evidence for CYP-mediated bioactivation, albeit with lower activity than vertebrates, but this issue needs yet much research. Through novel molecular tools, toxicology-relevant genes and proteins are being unraveled, from metallothioneins to heat-shock proteins and phase II conjugation enzymes, which highlights the importance of increasing genomic annotation as paramount to understand toxicant-specific pathways. However, little is known on how organic toxicants are stored, metabolized and eliminated, albeit some evidence from biomarker approaches, particularly those related to oxidative stress, suggesting that these molluscs' digestive gland is indeed responsive to chemical aggression. Additionally, cause-effect relationships between pollutants and toxicopathic effects are little understood, thus compromising, if not the deployment of these organisms for biomonitoring, at least understanding how they are affected by anthropogenically-induced global change.

#### Edited by:

Giovanna Ponte, CephRes and SZN, Italy

#### Reviewed by:

Paco Bustamante, University of La Rochelle, France Fuencisla San Juan, University of Vigo, Spain Carlos Rosas, Universidad Nacional Aitonoma De Mexico, Mexico

> \*Correspondence: Pedro M. Costa<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Invertebrate Physiology, a section of the journal Frontiers in Physiology

Received: 01 February 2017 Accepted: 03 April 2017 Published: 20 April 2017

#### Citation:

Rodrigo AP and Costa PM (2017) The Role of the Cephalopod Digestive Gland in the Storage and Detoxification of Marine Pollutants. Front. Physiol. 8:232. doi: 10.3389/fphys.2017.00232 Keywords: aquatic toxicology, mollusca, Cephalopoda, biomarkers, toxicological pathways, bioaccumulation

#### INTRODUCTION

Cephalopods are a particular group of invertebrates that share many important features with highorder animals as a result of convergent evolution, with emphasis on nervous system function. These features are, nonetheless, analogs to those of chordates, as cephalopods hold the basic molluscan body plan. Molluscs, however, form a cunningly diverse group of animals, ranging from sedentary filter feeders like bivalves to the giant predator squid Architeuthis. Among predators, cephalopods are of special interest in terms of anthropogenic impacts onto food webs, as they feed on a wide range of live prey and have high growth and metabolic rates (see Mangold, 1983), which poses important questions regarding bioaccumulation and tolerance to chemical

**152**

stressors. However, the physiological and molecular mechanisms underlying toxicopathological effects and detoxification processes in cephalopods are not well understood, albeit the importance of other molluscs, especially bivalves, in biomonitoring and substance testing. Overall, most studies concerning exposure of cephalopods to marine contaminants relate to the accumulation of trace elements, in most cases being limited to commercial species (see Penicaud et al., 2017).

It is suggested that storing metals in various tissues can be an important strategy to cope with metal toxicity (see Miramand and Bentley, 1992; Bustamante et al., 2000; Raimundo et al., 2005 plus the recent review by Penicaud et al., 2017). This efficient strategy, which likely minimizes energetic costs, is seemingly common among Nautiloid and Coleoid cephalopods (Bustamante et al., 2000). However, the mechanisms involved even in these basic processes are not fully resolved, particularly in the case of organic contaminants. This gap noticeable for invertebrates in general, bivalves included, albeit the attention the digestive gland has been receiving as target for bioaccumulation and biomarker analysis for being the analog the vertebrate liver. While there is indication that detoxification and excretion processes may indeed occur with the assistance of specialized cells (Costa et al., 2014), this issue is not entirely consensual as the digestive gland is able to provide long-term storage of both toxic and essential metals, such as Cd and Zn (Bustamante et al., 2002b) The growing level of genomic annotation for bivalves and a few cephalopods also indicates that CYP-like enzymes and respective organic xenobiotic pathways are active in molluscs (Cheah et al., 1995).

As such, the present review aims at summarizing the stateof-the-art on the role of digestive gland in the detoxification of organic and inorganic contaminants in cephalopods, emphasizing the comparative microanatomy, physiology, and molecular processes among various groups of molluscs that, however, indicate that toxicological pathways in cephalopods may be more diverse and complex than anticipated.

# FORM AND FUNCTION OF THE CEPHALOPOD DIGESTIVE GLAND

The molluscan digestive gland is a multi-task annex to the digestive tract, involved in secretion of digestive enzymes, extra and intracellular digestion, substance storage and excretion (Bidder, 1966). As in many invertebrates, organs are called to perform multiple functions due to reduced differentiation comparatively to vertebrates. The basic structure of the digestive gland is well-conserved among molluscs, being formed by blindend indigitations called "tubules" or more accurately, diverticula, being connected to the gut (specifically to the caecum, in cephalopods) by ducts (Budelmann et al., 1997). Refer to **Figure 1** for a comparative overview of the molluscan digestive gland. In cephalopods, albeit the lack of an absolute consensus, three distinct cells types have been identified. Digestive cells are the most abundant, followed by basal (replacement) and the more elusive excretory cells, characterized by a single, large hydropic vacuole that may bear mineral precipitates. Note that basal cells are commonly termed crypt and basophilic cells in gastropods and bivalves, respectively. Although demonstrated in cephalopods (e.g., Boucaud-Camou, 1968; Costa et al., 2014), the existence of specialized excretory cells is not consensual in other molluscs and their specific function in cephalopods is not well understood. There are, nonetheless, reports on changes in size and number of hydropic vacuoles of digestive gland cells of gastropods and bivalves as a result of exposure to mixed metallic and organic toxicants (Zaldidar et al., 2007; Lobo et al., 2010).

Cephalopod digestive gland epithelia are more complex than other molluscs' with respect to specialized endosomes. In fact, structures such as "boules" (vacuoles involved in digestion and enzyme secretion) and "brown bodies" (excretion of crystalline salts and amorphous materials) are seemingly exclusive. The lack of detailed studies integrating digestive gland histology and cytology with molecular pathways, as well as the lack of comparative studies between molluscan taxa hinders understanding how molluscs evolved to handle hazardous substances.

Accumulation of toxicants in the digestive gland depends on their mechanisms of apical entry. Bustamante et al. (2002b) revealed that Cd and Zn enter the cephalopods' digestive gland directly via food and indirectly via blood, in the latter case if uptake occurs from seawater. Nonetheless, the same authors disclosed that the elimination of these metals is faster if uptaken through water. It must be noted that the existence of a closed circulatory system in these molluscs, likely render the organ particularly efficient for nutrient absorption and as a filtering system for peripheral fluids. Indeed, unlike bivalves for instance, the cephalopod digestive gland possesses an intricate network of arteriole-like blood vessels (e.g., Swift et al., 2005; Costa et al., 2014). To these features is added the ability to form (and eventually release) mineral corpuscles called spherulae (spherocrystals) in the basal cells, first noticed by Martoja and Marcaillou (1993) and more recently described by Costa et al. (2014) which may thus have an important role in metal homeostasis, similarly to what has been suggested for some gastropods (Volland et al., 2012).

The first descriptions of the microstructure of the cephalopod digestive gland are almost as old as histology itself (refer to the pioneer works by Frenzel, 1886 and Cuénot, 1907). These were complemented by important histochemical descriptions being made from the 1960s onward that favored structural and digestion-related aspects (e.g., Boucaud-Camou, 1968; Semmens, 2002; Martinez et al., 2011). However, even for bivalves, which are the most investigated invertebrates by toxicologists, there are many gaps about the relation between form and function of the digestive gland and toxicant metabolism.

# THE DIGESTIVE GLAND IN BIOACCUMULATION AND DETOXIFICATION OF METALS

The vast majority of literature focusing on toxicants in cephalopods relates to metals, as cephalopods are known to bioaccumulate impressive amounts of hazardous elements, like Cd, albeit others, such as Hg, appearing to be less significant,

FIGURE 1 | Comparative histology of the molluscan digestive gland (paraffin sections). bc, basal cells (also called replacement; crypt, basophilic or pyramid cells); dc, digestive cells; dv, digestive vacuoles; hm, haemocytes; it, intertubular tissue; tl, tubule lumen. (A) Digestive gland of the common octopus (Octopus vulgaris) evidencing large digestive tubules (diverticula) formed mostly by digestive cells. The distinctive digestive vacuoles of cephalopods are naturally pigmented and traditionally referred by the French term "boules." Haematoxylin & Eosin. Scale bar: 25 µm. (B) Micrograph of the digestive gland of a cuttlefish (Sepia officinalis), showing a similar structure to that of Octopus. Brown bodies (bb) are distinctive of sepioids, being comprised of amorphous, undigested, materials. Tetrachrome stain. Scale bar: 25 µm. Inset: Basal cells were observed to hold calcic spherulae that include other metals as well, embedded in a proteinaceous matrix but the issue needs further research. The presence of calcium in spherulae in basal cells is here determined histochemically (stained black) through the von Kossa reaction, counterstained with Nuclear Fast Red (arrowhead). (C) Section through the digestive gland of the marine gastropod Onchidella celtica (Pulmonata), evidencing a similar structure and to that of cephalopods, albeit differences in the histochemical signal of digestive vacuoles, here predominantly blueish (from sugars), likely due to the herbivore feeding regime. The staining is similar to that of the preceding panel. The specimen was fixated in Zenker's solution, which contains (potassium) bichromate that reacts with metallic compounds originating yellow-orange deposits (arrowheads), once again visible in basal cells. Scale bar: 25 µm. (D) Section across the digestive gland of a bivalve (Ruditapes decussata), stained with Haematoxylin and Eosin. The tubules are smaller than previous examples and digestive cells less intricate with respective to variety, quantity and natural coloration of digestive vacuoles, regardless of digestive phase (which is similar among all panels). Basal cells are again evident and bear vesicular-like structures, potentially spherulae or similar. Note the wider and sparser intertubular tissue within which haemocytes can be found, as bivalves have an open circulatory system. Scale bar: 12 µm.

which indicates metal- and organ-specific pathways (Penicaud et al., 2017). It has been shown that the digestive gland holds the highest concentrations of essential (like Cu and Zn) and non-essential (such as Ag, Cd and Pb) metals in several cephalopods, with emphasis on Sepia and Octopus, comparatively to other organs, mantle and arms included, the latter of which raise particular concerns regarding human consumption (e.g., Raimundo et al., 2004, 2005; Seixas et al., 2005; Bustamante et al., 2006; Pereira et al., 2009). However, the digestive gland has been identified as the main metal accumulation organ even in the Nautiloidea (Pernice et al., 2009). Available data suggests that the cephalopod digestive gland is particularly efficient in the retention of these elements, likely as a function of chelating agents, especially proteins. In fact, there have been a few works that related concentrations of metals in this organ with different types of largely undisclosed proteins distributed through several subcellular partitions. For instance, Raimundo et al. (2010a) noticed that, in the common octopus, Pb was associated to unknown high molecular weight proteins while Zn, Cu and Cd showed high affinity to both high and low molecular weight proteins. Somewhat similar associations were recorded in the digestive glands of red arrow squid (Nototodarus gouldi) by Finger and Smith (1987) and in the common cuttlefish by Bustamante et al. (2006). These authors suggested that these associations may relate to different metal detoxification mechanisms, in many cases likely modulated by metal burden per se, through the induction of chelating proteins. Interestingly, Costa et al. (2014), demonstrated histochemically that the metalcontaining spherulae in cuttlefish digestive gland basal cells are formed by a matrix of proteinaceous materials, being released into the lumen of tubules as cells differentiate. However, there seems to be some selectivity in the accumulation of metals in spherulae. Unlike Cu, which appears to be accumulated mostly in these structures, Fe accumulates essentially in the cytosol of digestive cells, which has been confirmed histochemically in the independent works by Martoja and Marcaillou (1993) and Costa et al. (2014) with Sepia officinalis. Altogether, basal cell spherulae may be common in molluscan digestive glands, even though their role and formation is not well understood. Volland et al. (2012), for instance, demonstrated the existence of these structures in basal cells in the marine gastropod Strombus, albeit being seemingly not involved in Cd bioaccumulation upon induced exposure. In agreement, Bustamante et al. (2002a) reported that, in several different cephalopods, Cd is mostly cytosolic.

It has also been suggested that metallothioneins (MTs), well described in bivalves, play an important role in the formation of spherulae (Martoja and Marcaillou, 1993). However, in cephalopods, MT induction in the digestive gland may not be entirely consistent with exposure to metals, even in the case of exposure to strong inducers such as Cd (Bustamante et al., 2002a; Raimundo et al., 2010b; Rodrigo et al., 2013). It is possible that elements such as Cd, are not involved in detoxification via spherulae (thus remaining in the cytoplasmic fraction), which in its turn, relies on MT and, potentially, unknown high molecular weight proteins (see Penicaud et al., 2017, for a summary). This interesting perspective, indicates that there can be novel mechanisms of toxicity and detoxification of non-essential metals like Cd in cephalopods that need to be unraveled. As such, the known mechanisms for MT expression, described essentially for vertebrates, namely those relying on metallothionein transcription factor (MTF) mediation may not apply entirely or at all. In **Table 1** are summarized the most relevant among the very few publications on biomarker responses in cephalopod digestive gland in an ecotoxicological context, which includes also the MT response as potential indicator of exposure to metals.

# EVIDENCE FOR THE METABOLISM OF ORGANIC TOXICANTS

Although far less common than for metals, some studies addressed the issue of bioaccumulation of various organic hazardous substances in the cephalopod digestive gland, from polycyclic aromatic hydrocarbons (PAHs) and polychlorinated biphenyls (PCBs) to amnesic shellfish toxin (Costa et al., 2005, 2009; Danis et al., 2005; Storelli et al., 2006; Semedo et al., 2014). Nevertheless, the pathways of detoxification and elimination of organic substances, pollutants and toxins (endogenous or exogenous) that are called bioactive compounds are strikingly more complicated than metal chelation and expression of chelators. These mechanisms have been described mostly for mammals and are not consensual for invertebrates, once again indicating important knowledge gaps, in spite of the relevance of molluscs for ecotoxicologists.

The pathways for drug metabolism, meaning phases I (biotransformation) and II (conjugation), to which is now added a phase III (elimination) have been described in the vertebrate liver (see Ferreira et al., 2014, and references therein). Their functioning in invertebrates is not entirely consensual, in part due to the differential response of common biomarkers between vertebrates and invertebrates. For instance, one of the most important systems involved in phase I, the cytochrome P450 (CYP) monooxygenase complex (involved in the detoxification of many bioactive xenobiotics), has long been shown to hold similarities between invertebrates and vertebrates using cDNA probes and Western blotting techniques (Livingstone, 1994). This includes evidence that at least some octopodid cephalopods express CYP isoenzymes, albeit at reduced levels comparatively to vertebrates (Cheah et al., 1995). The CYP systems are localized mainly in the microsomes of the digestive gland cells of molluscs, although it was also found in other tissues like gills and even in haemocytes (Oehlmann and Schulte-Oehlmann, 2003). Still, the relevance of CYP1A in molluscs (involved in the metabolism of important PAHs, polychlorinated biphenyls and dioxins, as examples) has been disputed through works with bivalves, in favor of other forms, such as CYP4 (Chaty et al., 2004). In contradiction, there is some evidence for CYP1A activity and induction in terrestrial molluscs exposed to toxicants (Snyder, 2000, for a review). It is also important to mention that the pathways leading to increased CYP expression, usually involving xenobiotic-activated nuclear receptors (XANRs), are not well understood in invertebrates (see Richter and Fidler, 2014). In fact, the cephalopod equivalent for the aryl hydrocarbon receptor (Ahr) pathway, which is responsible for increased expression of CYP1A by PAHs and similar in vertebrates, remains a mystery. Still, there are promising findings with bivalves regarding its expression in gills and digestive glands exposed to Ahr agonists like benzo[a]pyrene (e.g., Châtel et al., 2012).

Schlenk and Buhler (1988) suggested that CYPs are more important in the metabolism of endogenous substrates rather than xenobiotic metabolism in Polyplacophora digestive glands, whilst other works evidenced some induction in bivalves (concerning enzyme content and activity) by agents ranging from pharmaceuticals to acrylamide, although unable to metabolize compounds that can be biotransformed in vertebrates or induced in a reduced extent (see for instance Galli et al., 1988; Larguinho et al., 2014). Also, Cheah et al. (1995) disclosed modest induction of CYPs in Octupus pallidus digestive gland after exposure to known inducers like β-naphtoflavone and Aroclor, with increased activity of ethoxycoumarin-O-deethylase (ECOD) but not for ethoxyresorufin O-deethylase (EROD), which is one of the best accepted biomarkers of exposure to bioactive pollutants in vertebrates. This finding is in agreement with the work by Schlenk and Buhler (1988) with chitons, even though Semedo et al. (2014) found negligible activities of both enzymes in wild Octopus vulgaris.

The activity of phase II enzymes, such as glutathione Stransferase or UDP-glucosyltransferase have been found in cephalopod digestive gland, with evidence that they can be induced by environmental toxicants but their specificity toward a specific class of substances has yet to be demonstrated (Schlenk and Buhler, 1988; Rodrigo et al., 2013; Le Pabic et al., 2015), since phase II enzymes can be induced by many factors, oxidative stress in particular. Inclusively, they were found to be promising targets in Octopus paralarvae exposed to metals (Nicosia et al., 2015). Altogether, less specific biomarkers, therefore responsive unspecific physiological challenge, yield more promising outcomes in the cephalopod digestive gland, among which are included superoxide dismutase (SOD) and catalase activities or glutathione induction (Semedo et al., 2012; Rodrigo et al., 2013; Nicosia et al., 2015). Note, though, that there are indications that the activity of oxidative stress enzymes can be significantly modulated by age in cephalopods (Zielinski and Pörtner, 2000). Interestingly, Oellermann et al. (2012) found that cuttlefish may adjust cardiac mitochondrial metabolism per se to adjust to thermal challenge, which indicates the ability to fine tune mitochondria-mediated oxidative functions under stress. This may explain the relevance of oxidative stress biomarkers in these animals, despite the missing link between mitochondrial function and toxicant-induced oxidative stress.

Another interesting target could be heat-shock proteins (HSPs) and similar chaperones that interfere with gene expression pathways. They have been described at the transcriptome level in several molluscs, including a few cephalopods (reviewed by Wang et al., 2013), and can be pointed as promising biomarkers (refer also to Oehlmann and Schulte-Oehlmann, 2003). In spite of the absence of similar work on cephalopods, with the exception of the work by Nicosia et al. (2015) wit h Octopus paralarvae (whose transcript levels increased with Cd exposure concentration), HSP70 induction has provided interesting results in other molluscs (bivalves in particular) subjected to several environmental stressors, from bacterial infection to metals, PAHs, pharmaceuticals, and biotoxins (Köhler et al., 1996; Boutet et al., 2003; Cellura et al., 2006; Song et al., 2006; Mello et al., 2012; Gust et al., 2013).

Altogether, detoxification of organic toxicants in the cephalopod digestive gland does occur, albeit potentially better adapted to dispose of toxins and unwanted by-products from feed and basal metabolism. It must also be considered that endpoints that are common biomarkers in vertebrates may not be the most adequate in molluscs, with particular respect to phase I. At the present, responses related to phase II and oxidative stress appear to be more viable biomarkers of exposure to bioactive organic toxicants, together with ECOD activity (refer to **Table 1**). Phase III of detoxification is still a subject of debate, being essentially reported for vertebrates, fish included. The action of specific transporters, like ATP-binding cassette (ABC) transporters, located in digestive tract cells, is described for the elimination of xenobiotic metabolites (reviewed by Ferreira et al., 2014). Even though not yet scrutinized in cephalopods, ABC transporters have been investigated in other molluscs and their action in gills during metal-induced challenge has already been reported (e.g., Della Torre et al., 2015). It would be of relevance

TABLE

1

Continued

to attempt to relate form and function of excretory vacuoles in digestive gland cells with phase III activity. Additionally, while the lack of genomic annotation is a clear drawback to unravel potentially novel mechanisms of detoxification through phases I to III in cephalopods, with the advent of state-of-the-art proteomics, metabolomics, (epi)genomics and transcriptomics tools (next-generation sequencing of genomic DNAs and RNAs), plus bioinformatics, more mechanistic information could be retrieved regarding detoxification pathways in cephalopods and other molluscs while assisting biomarker discovery.

#### CONCLUDING REMARKS

Even though it remains to be seen whether cephalopods can have such a significant role in biomonitoring as bivalves, the importance of these animals in ecosystem functioning, fisheries and more recently aquaculture dictates the relevance of investigating how these animals cope with anthropogenic pressures than endanger their habitats. The still scarce literature on physiological and molecular pathways related to detoxification of noxious substances suggests, however, that the digestive gland plays a major role, much in similarity with the vertebrate liver. As genomic annotation for these animals is slowly coupled with traditional biomarkers approaches that until recently were considered more or less exclusive to vertebrates,

#### REFERENCES

it may now be inferred that the toxicological pathways in cephalopods are more complex than expected for both metallic and organic xenobiotics, steering toward a new direction to understand the adaptative mechanisms of cephalopods to impacted marine ecosystems.

#### AUTHOR CONTRIBUTIONS

AR and PC are responsible for the conceptual design and writing of the manuscript. PC supervised the work.

#### ACKNOWLEDGMENTS

Part of this work was supported by the COST Action FA1301 "A network for improvement of cephalopod welfare and husbandry in research, aquaculture and fisheries (CephsInAction)". The authors are thankful to J. Raimundo (IPMA), S. Caeiro (CENSE), and M.H. Costa (MARE) for their support. The Portuguese Foundation for Science and Technology (FCT) is acknowledged for the funding for MARE through the strategic programme UID/MAR/04292/2013, plus the grants SFRH/BD/109462/2015 to AR and IF/00265/2015 to PC. The research project GreenTech (PTDC/MAR-BIO/0113/2014), also funded by FCT, is acknowledged as well.

heat shock and Vibrio anguillarum, but not by V. splendidus or Micrococcus lysodeikticus. Dev. Comp. Immunol. 30, 984–997. doi: 10.1016/j.dci.2005.12.009

Physiol. B Biochem. Mol. Biol. 125, 147–160. doi: 10.1016/S0305-0491(99) 00162-5

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2017 Rodrigo and Costa. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Hypoxic Induced Decrease in Oxygen Consumption in Cuttlefish (Sepia officinalis) Is Associated with Minor Increases in Mantle Octopine but No Changes in Markers of Protein Turnover

Juan C. Capaz 1 †, Louise Tunnah2 †, Tyson J. MacCormack <sup>2</sup> , Simon G. Lamarre<sup>3</sup> , Antonio V. Sykes <sup>1</sup> \* and William R. Driedzic<sup>4</sup> \*

#### Edited by:

Graziano Fiorito, Stazione Zoologica Anton Dohrn, Italy

#### Reviewed by:

Claudio Agnisola, University of Naples Federico II, Italy Panagiotis Grigoriou, Hellenic Centre for Marine Research, Greece

#### \*Correspondence:

Antonio V. Sykes<EMAIL_ADDRESS>William R. Driedzic<EMAIL_ADDRESS>
† These authors have contributed equally to this work.

#### Specialty section:

This article was submitted to Invertebrate Physiology, a section of the journal Frontiers in Physiology

Received: 05 January 2017 Accepted: 11 May 2017 Published: 26 May 2017

#### Citation:

Capaz JC, Tunnah L, MacCormack TJ, Lamarre SG, Sykes AV and Driedzic WR (2017) Hypoxic Induced Decrease in Oxygen Consumption in Cuttlefish (Sepia officinalis) Is Associated with Minor Increases in Mantle Octopine but No Changes in Markers of Protein Turnover. Front. Physiol. 8:344. doi: 10.3389/fphys.2017.00344 <sup>1</sup> Centro de Ciências do Mar do Algarve, Universidade do Algarve, Faro, Portugal, <sup>2</sup> Department of Chemistry and Biochemistry, Mount Allison University, Sackville, NB, Canada, <sup>3</sup> Département de Biologie, Université de Moncton, Moncton, NB, Canada, <sup>4</sup> Department of Ocean Sciences, Memorial University of Newfoundland, St. John's, NL, Canada

The common cuttlefish (Sepia officinalis), a dominant species in the north-east Atlantic ocean and Mediterranean Sea, is potentially subject to hypoxic conditions due to eutrophication of coastal waters and intensive aquaculture. Here we initiate studies on the biochemical response to an anticipated level of hypoxia. Cuttlefish challenged for 1 h at an oxygen level of 50% dissolved oxygen saturation showed a decrease in oxygen consumption of 37% associated with an 85% increase in ventilation rate. Octopine levels were increased to a small but significant level in mantle, whereas there was no change in gill or heart. There were no changes in mantle free glucose or glycogen levels. Similarly, the hypoxic period did not result in changes in HSP70 or polyubiquinated protein levels in mantle, gill, or heart. As such, it appears that although there was a decrease in metabolic rate there was only a minor increase in anaerobic metabolism as evidenced by octopine accumulation and no biochemical changes that are hallmarks of alterations in protein trafficking. Experiments with isolated preparations of mantle, gill, and heart revealed that pharmacological inhibition of protein synthesis could decrease oxygen consumption by 32 to 42% or Na+/K<sup>+</sup> ATPase activity by 24 to 54% dependent upon tissue type. We propose that the decrease in whole animal oxygen consumption was potentially the result of controlled decreases in the energy demanding processes of both protein synthesis and Na+/K+ ATPase activity.

Keywords: European cuttlefish, Sepia officinalis, HSP70, octopine, polyubiquitinated protein, ventilation frequency

# INTRODUCTION

The European cuttlefish (Sepia officinalis) is a dominant species in the north-east Atlantic Ocean and Mediterranean Sea. It is a benthic species that occurs to depths of 200 m. Sexually mature and juvenile individuals migrate to coastal grounds in the spring; the latter in pursuit of abundant food sources (Pierce et al., 2008; Bloor et al., 2013). These geographic areas are at risk of becoming

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hypoxic because of the increase of nutrients and organic matter into the coastal waters (Van Den Thillart et al., 1994; Grantham et al., 2004; Chan et al., 2008). Hypoxic conditions occur naturally in nature (Ekau et al., 2010); however, there are reports of a global oceanic oxygen decline (Schmidtko et al., 2017), which is becoming a problem for marine biodiversity (Vaquer-Sunyer and Duarte, 2008) and an important environment stressor to marine species (Cosme and Hauschild, 2016; Townhill et al., 2017). Hypoxia is also a potential challenge under intensive aquaculture conditions (Brett, 1979; Delaney and Klesius, 2004; Burt et al., 2013) due to the high density of animals and because of the costs associated with keeping oxygen saturation at normal levels. Routine oxygen consumptions of sepioids are within the same range of those of octopus species (Boyle, 1991) and optimal oxygen saturations for Octopus vulgaris are reported to be in between 100 and 65% dissolved oxygen saturation (DO2) for proper food intake and growth, while suboptimal (modest hypoxia) values range from 65 to 35% DO<sup>2</sup> (Cerezo Valverde and García García, 2005). These suboptimal values may be reached with a combination of high densities and temperatures in Mediterranean countries, where cuttlefish culture will take place. This is relevant because oxygen availability limits thermal tolerance in S. officinalis (Melzner et al., 2006), O<sup>2</sup> consumption rates rise with increasing temperature up to a metabolic threshold (Melzner et al., 2007) related to the oxygen-binding properties of haemocyanin, which lowers with increasing temperature (Brix et al., 1994). As such, it is important to have an understanding of the response of cuttlefish to challenges of a suboptimal oxygen environment that is considered to be either ecologically and/or practically relevant.

S. officinalis, as with other cephalopods, exhibit a decrease in oxygen consumption below a critical environmental oxygen level (Houlihan et al., 1982; Johansen et al., 1982; De Wachter et al., 1988; Cerezo Valverde and García García, 2005; Seibel et al., 2014). Severe hypoxia induces a mobilization of mantle glycogen pools in this species with concomitant accumulation of the anaerobic end product octopine (Storey and Storey, 1979; Storey et al., 1979). The production of octopine is catalyzed by the enzyme octopine dehydrogenase (ODH) that occurs in particularly high levels in mantle, ventricle, and tentacle relative to other tissues (Storey, 1977). Despite this existing information, the biochemical responses of this species, at the tissue level, to environmentally relevant levels of hypoxia have not been fully characterized, so the importance of anaerobic octopine production under these conditions is unclear. In addition to octopine accumulation, more recent studies with the exceptionally hypoxia tolerant Humboldt (jumbo) squid, Dosidicus gigas, reveal other aspects of the response to low oxygen levels that include a decrease in mantle contraction frequency (i.e., gill ventilation) and increases in mantle heat shock protein 70 (HSP70) and ubiquinated proteins (Trübenbach et al., 2013a,b, 2014). Here we question if these responses occur in S. officinalis as well, as it is considered a hypoxia-sensitive species (Storey et al., 1979).

Decreases in oxygen consumption that are not due to reduced spontaneous swimming activity must be related to decreases in other components of energy demand. Aside from muscle contraction, the two most energy demanding processes in cells are the maintenance of ionic gradients via Na+/K<sup>+</sup> ATPase and protein turnover (synthesis and degradation) (Wieser and Krumschnabel, 2001). The rapid growth rates of cephalopods have been suggested to occur due to high rates of protein synthesis and high efficiency of protein retention, which implies low protein degradation (Houlihan et al., 1990; Carter et al., 2009; Moltschaniwskyj and Carter, 2010). It is possible to measure the relationship between these processes and oxygen consumption by pharmacological inhibition of Na+/K<sup>+</sup> ATPase with ouabain and protein synthesis with cycloheximide (Wieser and Krumschnabel, 2001; Agin et al., 2003). Here we use these pharmacological tools on isolated preparations to assess the potential tissue specific contribution of Na+/K<sup>+</sup> ATPase activity and protein turnover to the whole animal decrease in oxygen consumption.

Mantle is the largest tissue of S. officinalis, corresponding to approximately 30–40% of body mass (Castro et al., 1992; Speers-Roesch et al., 2016). We focused our attention on this tissue for assessment of biochemical changes and to determine its potential contribution to alterations in metabolic demand under hypoxia. Although gill and heart represent only about 2 and 0.1% of body mass, respectively (Speers-Roesch et al., 2016), these tissues were also included in our study as they are essential organs to supply oxygen to the rest of the animal, which must maintain function under hypoxia, and have been previously connected to high fractional rates of protein synthesis (Houlihan et al., 1990).

## MATERIALS AND METHODS

# Ethical Statement

All the procedures were approved by CCMAR Animal Welfare Committee (ORBEA CCMAR-CBMR) and Direcção-Geral de Alimentação e Veterinária (DGAV) of the Portuguese Government, according to National (Decreto-Lei 113/2013) and EU legislation (Directive 2010/63/EU) on the protection of animals used for scientific purposes. In addition, protocols were approved by institutional Animal Care Committees at each of the Canadian Universities where authors of this paper are based. Procedures were only applied to live animals by authorized users.

#### Animals

Cuttlefish (S. officinalis) were reared according to the latest culture technology described in Sykes et al. (2014). All experiments used juveniles produced from eggs laid by a F6 captive stock. The mass of group 1 was 59.3 ± 5.4 g (N = 18) and of group 2, used only in mantle oxygen consumption experiments to avoid limiting oxygen diffusion across thick mantle tissue, was 1.90 ± 0.14 g (N = 7). Experiments were done during May 2016, at CCMAR's Ramalhete Aquaculture Station (Ria Formosa, Portugal—37◦ 00′ 22.39′′N; 7◦ 58′ 02.69′′W). Temperature, salinity, and DO<sup>2</sup> were measured daily, at 9 h 30, in the stock tank. Both temperature and DO<sup>2</sup> were measured with a VWR DO220 probe, while salinity was measured with a VWR EC300 salinity meter. Water temperature was 20.5 ± 1.27 (S.D.)◦C, salinity was 35.9 ± 0.9 (S.D.) g L−<sup>1</sup> and DO<sup>2</sup> level

was 101.1 ± 1.2 (S.D.)%. Cuttlefish were fed frozen grass shrimp (Palaemonetes varians) ad libitum on a daily basis.

#### Oxygen Uptake and Ventilation Frequency in Whole Animals

Two closed respirometry (Lefevre et al., 2016) acrylic chambers (48.5 × 28.5 × 29 cm), each with an external recirculating pump (Maxi-Jet PH MP400) and a PASCO PasPort PPS-2169 multiparameter sensor were used. The recirculating system had a total volume of 41 L. Seawater entered and exited on opposite sides of the chamber at a flow of 200 L h−<sup>1</sup> . Dissolved oxygen saturation data were collected in the upper right section of the chamber at 1 Hz in the PASCO Capstone software and at a temperature of 19.9 ± 1.36 (S.D.) ◦C. The chamber was completely isolated from any visual stimuli (both researchers and light) by the application of matte black stickers on the outer walls, except from above and one of the 48.5 cm sides. This side had a rectangular LED stripe (50 × 30 × 50 × 30 cm) attached that emitted red light (wavelength peak of 630 nm) until the chamber was completely filled with water and, thereafter, white light (wavelength peaks at 460, 520 and 630 nm) with an intensity of 80 lux. These conditions allowed for the recording of movement (from above) with a Logitech C615 HD camera, and of behavior and ventilation rates (from the side) with a GoPro Hero 3 <sup>+</sup> Black Edition. Video recordings were made for the duration of each experiment at 1,920 × 1,080 pixel resolution and 24 frames per s (fps), from above, and 1,920 × 720 pixel and 50 fps, from the side.

Initially, 6 animals were individually exposed to a DO<sup>2</sup> of 100% and 6 other animals to a DO<sup>2</sup> of 50% oxygen saturation for 60 min. Each condition was tested pairwise, 1 cuttlefish at 100% DO<sup>2</sup> vs. another at 50% DO2. At the end of this procedure all the animals were euthanized. Samples of mantle, gill, and heart were rapidly removed and frozen in liquid nitrogen for biochemical analysis. Rates of oxygen consumption MO˙ <sup>2</sup> were determined for animals exposed to 50% DO<sup>2</sup> during 60 min. In addition, 6 further control animals (DO<sup>2</sup> of 100%) were used to collect MO˙ <sup>2</sup> during 90 min, to increase the number of available data points for regression fitting. Background MO˙ <sup>2</sup> (resulting from consumption of bacteria and electrode) was also assessed with no animal (N = 3) in the chamber for 90 min and was found to be negligible.

Hypoxia at 50% DO<sup>2</sup> was achieved by gassing seawater with pure N<sup>2</sup> using a ceramic stone in a closed 90 L reservoir. This water was then injected into the chamber, until the 41 L respirometry recirculating system was filled, and where the animal was already resting. Animals were removed from the stock tank with a black net and transported in a 10 L black bucket filled with seawater to the weighing station. They were individually weighed in a black plastic tray and quickly transported to the chamber area of the experimental room which was illuminated with 1 lux red light conditions (Philips TL-D Colored 18W Red 1SL/25). An outer room containing the data collection stations was isolated from the holding chambers. To avoid inking in the experimental chambers, animals were left resting in the bucket for 5 min before being lightly sedated (mild anesthesia) by adding ethanol to a concentration of 2.5% (Fiorito et al., 2015). After sedation (≈2 min), animals were transferred to the chamber, which had 2 cm in height of seawater and thereafter filled with seawater from the 90 L reservoir (either 100 or 50% DO2). As soon as the chamber was filled (≈2 min) and air purged, the system was put in recirculating mode, light switched to white and data acquisition was started on all the probes. This filling time was enough for recovery from sedation at this concentration (Gonçalves et al., 2012).

Ventilation rates were determined from video recordings, according to the following sampling scheme for each animal: (a) first and last 10 min—10 samples of 1 min each; b) from 10 to 50 min in hypoxia and normoxia (control)—1 sample every 10 min. After the procedure, the chamber was emptied, the animal removed, and euthanized with 10% ethanol in seawater (Sykes et al., 2012), the brain was then bisected downwards and forwards, followed by two lateral cuts to sever the brain from the optical lobes (Lewbart and Mosley, 2012). No relevant movements in the chamber nor extreme behavior reactions (Gonçalves et al., 2012) were recorded in any video sampling.

#### Biochemical Assays

#### Octopine, Glucose, and Glycogen

Tissues were homogenized and then sonicated (QSonica q55) in 6% HClO<sup>4</sup> (mantle 1 g tissue: 9 mL HClO4; gill 1 g tissue: 4 mL HClO4), and subsequently centrifuged at 14,000 × g for 10 min at 4◦C. The supernatant was neutralized with 2 M KHCO3. Octopine was assayed in buffer containing 100 mM TRIS, 8 mM NAD+, and excess octopine dehydrogenase (ODH) at pH 9.3. The increase in absorbance at 340 nm, following addition of enzyme, was monitored in a microplate reader. Octopine was calculated based on the NADH extinction coefficient of 6.22 and the pathlength of the plate reader assay was experimentally determined in near IR using the absorbance of water at 975 nm according to the following equation (Lampinen et al., 2012).

$$\text{pathlength} = \frac{OD975\_{\text{assay buffer}} - OD900\_{\text{assay buffer}}}{0.173} \times 10 \, mm$$

ODH was isolated from frozen scallop, Placopecten magellanicus, adductor muscle by a modification of the method of Gäde (1985). Tissue was homogenized (25% w/v) in 100 mM TRIS (pH 7.5, 0.1 mM EDTA, 0.1 mM DTT) and centrifuged at 17,000 × g for 10 min at 4◦C. Ammonium sulfate was then added to the supernatant up to 65% saturation and stored at 4◦C for 7 days to precipitate ODH. The sample was centrifuged, the pellet was re-suspended in 100 mM TRIS (pH 7.5, 0.1 mM EDTA, 0.1 mM DTT), the sample was desalted on a Sephadex G-50 column, and fractions containing ODH activity were retained. Contaminating dehydrogenases were removed by passing the sample through a Cibachrome Blue F3GA agarose column (Affi-Gel Blue, Bio-Rad Laboratories, Hercules, California, USA). Fractions containing ODH activity were combined and subsequently desalted on a Sephadex G-25 column to ensure removal of all residual metabolites. The resulting sample yielded 2 bands on an SDS-PAGE gel, one at ∼43 kDa matching the molecular weight of ODH, and a second band of unknown identity at approximately

90 kDa. Lactate dehydrogenase activity was undetectable in the purified sample.

Glucose was analyzed in buffer containing 250 mM imidazole, 5 mM MgSO4, 10 mM ATP, 0.8 mM NADP+, excess glucose 6-phosphate dehydrogenase and thereafter treated with excess hexokinase. A standard curve was created for glucose analysis. Assays were conducted in a microplate reader at a wavelength of 340 nm. Glycogen was assessed as glucosyl units following amyloglucosidae treatment of the neutralized supernatant (Keppler and Decker, 1974). Free glucose was subtracted from the total glucosyl equivalents to yield glycogen levels. All organic reagents were purchased from Sigma-Aldrich.

#### Polyubiquitin and HSP70

Polyubiquitinated proteins and HSP70 were determined by western blots. Frozen tissues were sonicated in 9 vol of 50 mM TRIS buffer (pH 7.4) and then centrifuged for 5 min at 14,000 × g at 4◦C. The protein concentration in the supernatant was determined using the Bradford assay (Bio-Rad) and then adjusted to 3 µg/µL with the homogenization buffer. Tissue extracts were added to Laemmli sample buffer (1:1), boiled for 5 min and 4.5 µg of proteins were resolved by SDS-PAGE (TGX Stain-Free FastCast 10%, Bio-Rad). Resolved proteins were subsequently transferred to PVDF membranes, the membranes were blocked using 5% BSA before being incubated with the primary antibody. To analyze the levels of polyubquitinated proteins, we used an antibody that only recognized proteins containing K48-linked polyubiquitin chains (Abcam ab190061) but had no cross-reactivity to monoubiquitin or polyubiquitin of other linkages. Levels of HSP70 were quantified by loading known quantities of a recombinant HSP70 (Enzo Life Sciences, #ADI-SPP-758, standard levels of 15, 30, and 60 ng per lane) along with the samples and using an HSP70 antibody (#AS05 083A, Agrisera). In all cases, secondary detection was done using an anti-rabbit HRP-linked antibody (for polyubiquitin: #7074, Cell Signaling Technology; for HSP70: SAB-300, Enzo Life Sciences). The chemiluminescence signal was detected using a CCD camera system (ImageQuant LAS 500, GE Life Sciences). Band intensity was quantified using ImageJ (imagej.nih.gov).

#### Oxygen Uptake by Isolated Preparations

Tissue MO˙ <sup>2</sup> was measured in 0.20 µm filtered seawater containing an additional 10 mM KCl, 200 mM taurine, and 1 mM glucose. The concentration of taurine used here is representative of measured tissue taurine levels in this species (Maccormack et al., 2016). Cuttlefish were euthanized as described above and 5–10 mg samples of gill, systemic heart, or mantle muscle were collected, weighed, and cut into 2– 3 mg pieces using a razor blade. Tissue preparations from the same animal were then transferred to paired respirometry chambers (OX1LP, Qubit Systems Inc., Kingston, ON, Canada) containing 750 µL incubation medium maintained at 20 ± 0.1◦C with a recirculating refrigerated water bath. Chambers were calibrated daily and background O<sup>2</sup> consumption rates were negligible without tissue. Baseline MO˙ <sup>2</sup> was recorded for 300 s before addition of pharmacological agents and recordings were continued for an additional 300 s before experiments were terminated. Treatments consisted of 5 mM ouabain (e.g., Postel et al. (2000)) or 25 µM cycloheximide, a known antibiotic inhibitor of protein synthesis (Giuditta et al., 1968; Prozzo and Giuditta, 1973; Wieser and Krumschnabel, 2001) that has previously been applied in studies associated with learning in the species by blocking protein synthesis (Agin et al., 2003). Stock solutions were prepared in DMSO and delivered to the chambers in 10 µL. A tissue preparation from the same animal was run simultaneously with each drug treatment and exposed to 10 µL DMSO as a vehicle control. Chambers were thoroughly rinsed with 95% ethanol, ddH2O, and incubation medium between runs and treatments were alternated between respirometry chambers to control for potential chamber effects. Respirometry chambers were interfaced to a LabQuest data acquisition system and DO<sup>2</sup> readings were collected over 10 min using LoggerPro V3.8 software (Vernier Software and Technology, Beaverton, OR, USA). Initial tissue MO˙ <sup>2</sup> prior to treatment was determined from the linear decrease in DO<sup>2</sup> between 120 and 300 s and posttreatment MO˙ <sup>2</sup> was determined between 400 and 600 s. MO˙ 2 was linear with tissue mass (data not shown) and consistent with previous experiments (Maccormack et al., 2016).

## Data Analysis and Statistics

All values are expressed as mean ± s.e.m with the exception of water temperature, salinity, and DO<sup>2</sup> that are expressed as mean ± standard deviation. Statistical tests applied are stated in either the Results section or the legends to the figures. All data were tested for normal distribution with the Shapiro-Wilk test as well as for homogeneity of the variances with the Levene's test (Zar, 1999). Statistical difference was considered for P < 0.05.

# RESULTS

#### Whole Animal Experiment

**Figures 1A,B** show examples of dissolved oxygen variation over the sampling period for individual cuttlefish either exposed to 100 or 50% DO<sup>2</sup> over 90 or 60 min, respectively. Oxygen consumption by hypoxic animals was 37% lower than animals held at 100% air saturation (189 ± 23 vs. 119 ± 6 nmol/g min) (**Figure 1C**). Cuttlefish maintained under control normoxic conditions exhibited a decrease in ventilation rate over the 60 min holding period (y = −0.52x + 88) (**Figure 1D**). Animals held under hypoxia also displayed a decrease in ventilation rate but to a lesser extent (y = −0.1x + 102). After 60 min ventilation rate was 85% higher in hypoxic than normoxic individuals.

Octopine was significantly higher in mantle than in gill or heart regardless of oxygenation condition. Octopine levels were significantly higher in mantle of hypoxic than normoxic animals but there was no significant difference between levels in normoxic or hypoxic animals in either gill or heart. (**Figure 2A**). There was no significant difference in either free glucose or glycogen levels in mantle between normoxic or hypoxic held animals (**Figure 2B**). HSP 70 was significantly higher in gill than in mantle or heart regardless of oxygenation condition (**Figure 2C**). There was no significant difference between levels in normoxic or hypoxic animals in any of the three tissues. Similarly, there

exposed to 50% DO<sup>2</sup> (mild hypoxia). (C) Oxygen consumption expressed as nmol O2/(g wet weight animal) min. \* Indicates a statistically significant difference between normoxic and hypoxic conditions (2 tailed t-test; P = 0.025). N = 6 for normoxic and N = 5 for hypoxic conditions. (D) Ventilation rate expressed as cycles/min. Control, circles; Hypoxic, squares. N = 4 for all time points. The slopes are significantly different (P < 0.001).

was no significant difference in levels of polyubiquinated protein in mantle, gill or heart between treatment groups (**Figure 2D**).

#### Isolated Tissue Experiment

Pre-treatment oxygen consumption rates for gill, heart, and mantle are presented in **Figure 3A**. There was no significant difference in rates amongst the three tissues with values for all being approximately 500 nmol O2/gm wet weight. min. Rates for gill and heart are from animals weighing approximately 59 g; while those for mantle are from smaller/younger animals with a mass of approximately 2 g. Preparations were treated with cycloheximide and ouabain to assess how much of the decrease in oxygen consumption noted in the whole animal study could potentially be attributed to protein synthesis and Na<sup>+</sup> / K <sup>+</sup> ATPase, respectively. **Figure 3B** shows the percentage decrease in oxygen consumption for each of the tissues following correction for any DMSO effect. Cycloheximide had a substantial effect upon oxygen consumption with average decreases ranging from 32% for gill to 42% for mantle. Similarly, ouabain treatment resulted in average decreases in oxygen consumption from 24% for mantle to 54% for heart.

# DISCUSSION

#### Changes in MO˙ <sup>2</sup> and Ventilation Frequency under Hypoxia

Cuttlefish remained sedentary in the respirometry chambers under both normoxic and hypoxic conditions. As such, any difference in oxygen consumption between groups cannot be attributed to spontaneous swimming activity. When corrected for differences in temperature and body mass, the MO˙ <sup>2</sup> values reported here under control conditions are in the same range as rates previously noted (Johansen et al., 1982; Melzner et al., 2006, 2007; Grigoriou and Richardson, 2009; Lamarre et al., 2016). Here, exposure to 50% DO<sup>2</sup> for 60 min resulted in a 37% decrease in oxygen consumption. Although limited to 2 individuals of S. officinalis with a mean mass of 153 g, a decrease in MO˙ <sup>2</sup> of about 29% at a similar level of hypoxia was previously observed by De Wachter et al. (1988). Under comparable conditions, S. officinalis of body mass between 100 and 1,500 g showed a decrease in oxygen consumption of approximately 50% (Johansen et al., 1982). It is clear that a substantial reduction in MO˙ <sup>2</sup> is a common response to modest hypoxia

significance for octopine, HSP70, and polyubiquitinated proteins, was assessed with a 1-way ANOVA and for differences between glucose or glycogen levels with a t-test. N = 6 for all conditions except for free glucose in hypoxic mantle where N = 4. Differences between means or grouped means represent statistical difference (Tukey's multiple comparison test; P < 0.001). No differences were found in mantle free glucose and glycogen nor polyubiquitinated proteins (P > 0.05).

in S. officinalis; whether life could be sustained indefinitely at this level of hypoxia and if this will impact on the welfare (pain, suffering, distress, and lasting harm) of the animal remains unknown.

Ventilation rate under normoxic conditions decreased over the 60 min holding period in the same fashion as previously shown (Boal and Ni, 1996) and is presumably related to the stress of transferring the animals to the experimental chamber. After 60 min the ventilation rate recorded here was in the same range as previously noted in other studies (Bone et al., 1994; Boal and Ni, 1996; Boal and Golden, 1999; Melzner et al., 2006). Ventilation rate in hypoxic animals remained relatively stable with time and was 85% higher in hypoxic than normoxic cuttlefish after 60 min. To our knowledge this is the first report of rate of ventilation in hypoxic S. officinalis or any other cuttlefish species. The response differs from inactive, juvenile Humbolt squid where deep hypoxia resulted in a decrease in ventilation frequency, as well as contraction strength and ventilation volume per min (Trübenbach et al., 2013a). The mantle muscle of squid and cuttlefish consists of a main mass of circular fibers that are mitochondria-poor and primarily anaerobic, as well as inner and outer layers of mitochondriarich aerobic fibers (Bone et al., 1981; Mommsen et al., 1981). The central fibers are used in escape jetting contractions while the fibers of the inner and outer layers are responsible for rhythmical respiratory ventilation (Bone et al., 1981). It is probable that the increase in ventilation frequency observed here places additional demand to supply ATP to the contractile mechanisms under hypoxia even in the face of reduced oxygen consumption.

# Anaerobic Metabolism is Minimal

Octopine dehydrogenase is the dominant opine dehydrogenase in S. officinalis. We were unable to detect activity of alanopine, tauropine, or strombine dehydrogenase in this species' tissues (unpublished data). The level of ODH in mantle, heart and gill was 35, 20, and 2 µmol/min g wet weight, respectively (Speers-Roesch et al., 2016). The level of octopine in mantle was substantially higher than in heart or gill, consistent with the high activity of ODH in mantle and lower activity in the other two tissues.

Sixty min of 50% hypoxia was associated with an increase in mantle octopine of 2.5µmol/g. There was no change in octopine level in either gill or heart following the hypoxic period. The observed increase in octopine in mantle was low relative to the capacity to produce this metabolite. For instance, the increase in mantle octopine in S. officinalis forced to exercise and then subjected to hypoxia was 13µmol/g (Storey and Storey, 1979). Another study found a drastic elevation of octopine in the haemolymph of S. officinalis exposed to <30% O<sup>2</sup> hypoxia for about 60 min (Storey et al., 1979). A similar increase in mantle octopine occurred in hypoxic Humbolt squid (Seibel et al., 2014) and squid (Lolliguncula brevis) forced to swim to exhaustion (Finke et al., 1996). The modest increase in octopine observed in this study is in line with the lack of change in glycogen levels during the hypoxic challenge. Based on ATP equivalents that may be generated by aerobic and anaerobic metabolism, the contribution of octopine production to overall energy production is minimal.

The relative quantity of polyubiqutinated protein was similar in mantle, gill, and heart under control conditions. There was no impact of the hypoxic challenge in any of the tissues tested. This finding is in contrast to that for juvenile Humbolt squid where severe hypoxia resulted in a three-fold increase in ubiquitinated proteins with a size of approximately 100 kDa (Trübenbach et al., 2014). These authors proposed that the increase in polyubiquintinated protein in association with a decrease in levels of at least two abundant proteins in the same size range (HSP90 and alpha-actinin) provides protection under hypoxia via a number of mechanisms, including the provision of certain amino acids for anaerobic energy production (Trübenbach et al., 2014). In S. officinalis food deprivation results in almost total depletion of digestive gland triglyceride, MO˙ <sup>2</sup> decreases by approximately 30%, and mantle polyubiquitin mRNA and polyubiquitinated proteins increase, suggesting an increase in protein degradation via the ubiquitin-proteosome pathway to provide amino acids for energetic purposes (Lamarre et al., 2012, 2016). Although in this species there appear to be mechanisms to increase protein breakdown in association with decreased MO˙ 2, we found no evidence that this occurs under modest hypoxia, at least based on levels of polyubiquinated protein.

HSP70 is a well-recognized stress protein that plays a role in maintaining protein integrity via chaperoning newly synthesized and unfolding proteins (Balchin et al., 2016). HSP70 levels were significantly higher in gill than in mantle or heart and did not change in any tissue with exposure to hypoxia. Relatively high HSP70 levels in S. officinalis gill may be related to the high rate of protein synthesis in gill relative to mantle based on direct measurements (Lamarre et al., 2016), and relative to heart and mantle based on total RNA levels (Lamarre et al., 2012). Furthermore, since gill is at the interface with the aquatic environment it may experience more perturbations than other tissues and so has higher constitutive levels of HSPs as protection. HSP70 levels increased in mantle of hypoxic juvenile Humbolt squid (Trübenbach et al., 2013b) but not in adult animals (Seibel et al., 2014). Thus, the generality of a HSP70 response to hypoxia in cephalopods is unresolved but it is clear that under the conditions of the current study there was no change in HSP70 level.

The minimal increase in octopine content, along with stability in glycogen pools, polyubiquitinated protein level, and HSP70 levels in cuttlefish subject to 50% hypoxia for 1 h suggests that anaerobic metabolism is activated to only a limited extent and that the challenge is not particularly stressful to the animal. The 37% decrease in MO˙ <sup>2</sup> must be associated with an equivalent decrease in energy demand. Given that the animals are quiescent under both control and hypoxic conditions, the question becomes as to what metabolic processes are decreased to maintain energy balance.

# Potential Sites of Energy Reduction—Insights from Isolated Tissue Experiments

There was no difference in the rate of oxygen consumption between tissue slices of gill, heart, and mantle. The in vitro activity of citrate synthase, a qualitative indicator of aerobic metabolism, was approximately 20-fold higher in S. officinalis heart than in mantle or gill which were similar to one another (Speers-Roesch et al., 2016). The relatively low rate of oxygen consumption in heart noted here is likely due to the non-contractile nature of the preparation and reflects the basal metabolism of quiescent cells. The heart and gill samples were from juvenile animals and the mantle from hatchlings. Smaller animals have higher mass specific rates of oxygen consumption than larger animals (Johansen et al., 1982; Grigoriou and Richardson, 2009). It is likely that if oxygen consumption of mantle had been conducted on tissue removed from animals the same size as that used for gill and heart that the rate for mantle would have been relatively lower.

One of the major energy demanding processes in noncontracting cells is protein synthesis. In food deprived cuttlefish, both MO˙ <sup>2</sup> and protein synthesis are decreased in concert. A 36% decrease in whole animal MO˙ <sup>2</sup> was associated with a 63% decrease in fractional rate of protein synthesis in both mantle and gill (Lamarre et al., 2016). The importance of these findings in the context of the current study is that it demonstrates that protein synthesis is a regulatable element of the energy demand machinery in S. officinalis. Here we show that the inclusion of cycloheximide decreases MO˙ <sup>2</sup> in isolated gill, heart, and mantle preparations. Based on studies with goldfish and trout it is likely that the level of cycloheximide utilized here resulted in a total shut down of protein synthesis (Wieser and Krumschnabel, 2001), as previously suggest by the Agin et al. (2003) results on the species and because the amount used here was more than 3 times higher than the dosages used in Octopus vulgaris (Prozzo and Giuditta, 1973) and Loligo pealii (Giuditta et al., 1968), which resulted in 80% protein synthesis inhibition in the optic globe with 100 µg/mL and 95– 96% inhibition in squid axons with 200 µg/mL, respectively. If it is assumed that all tissues in the quiescent animals are consuming oxygen at similar rates, a total inhibition of all protein synthesis would be required to meet the whole animal decrease in MO˙ <sup>2</sup> of 37%. This is highly unlikely given that even in an anoxic resistant fish, oscar (Astronotus crassipinnis), protein synthesis is decreased by only 55% and 60–85% in muscle and liver, respectively, at a DO<sup>2</sup> of 10% (Lewis et al., 2007). Therefore, although a decrease in protein synthesis could potentially contribute to the decrease in whole animal MO˙ <sup>2</sup>, there must be additional mechanisms to reduce energy expenditure.

Na+/K<sup>+</sup> ATPase is a major driving force for many energy dependent ion transport processes and is considered one of the major cellular sites for ATP utilization.

Na<sup>+</sup> /K<sup>+</sup> ATPase has been identified in numerous tissues of S. officinalis (Donaubauer, 1981) and is presumably ubiquitous. A ouabain sensitive Na+/K<sup>+</sup> ATPase has been identified in gill of squid (Doritheutis plei) (Proverbio et al., 1988) and more recently, Na <sup>+</sup>/K<sup>+</sup> ATPase in Sepia gill was shown to increase in response to increases in water CO<sup>2</sup> level (Hu et al., 2011). In the hypoxia tolerant intertidal clam, Mercenaria mercenaria, a severe hypoxic exposure resulted in a decrease in the maximal in vitro enzyme activity of Na+/K<sup>+</sup> ATPase (Ivanina et al., 2016). As with protein synthesis, these studies illustrate that Na+/K<sup>+</sup> ATPase is a site that can be controlled to alter energy demand. Here it is shown with isolated tissues that inhibition of Na+/K<sup>+</sup> ATPase with ouabain decreased MO˙ <sup>2</sup> by ∼20% in mantle and 50–60% in gill and heart. Given that the Na+/K<sup>+</sup> ATPase is one of the largest consumers of ATP in cells, it is likely that a curtailment of this process occurs in S. officinalis under hypoxia.

# CONCLUSIONS

A hypoxic challenge to S. officinalis of 50% DO<sup>2</sup> results in only a minor activation of anaerobic metabolism as assessed by octopine accumulation. There were no changes in levels of HSP70 and polyubiquitinated proteins suggesting no alteration in rates of protein breakdown. The animal responds with an increase in ventilation and an overall decrease in metabolic rate, potentially due to decreases in both rates of protein synthesis and Na+/K<sup>+</sup> ATPase activity. The relatively high level of HSP70 in gill compared to mantle and heart is a novel finding and may be related to the high rates of protein synthesis in this tissue.

# AUTHOR CONTRIBUTIONS

SL, TM, AS, and WD participated in the conception and design of research; JC and AS performed the animal husbandry; all authors performed the experiments, acquired and analysed data of work; SL, TM, AS, and WD performed interpretation of data; all authors drafted, edited and approved the final version of the manuscript. All authors agree to be accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved.

# FUNDING

TM, SL, and WD were supported by Natural Sciences and Engineering Research Council of Canada (NSERC) Discovery grants. WRD holds the Canada Research Chair in Marine Bioscience. LT held a New Brunswick Innovation Foundation (NBIF) STEM Graduate scholarship. AVS was supported by Fundação para a Ciência e a Tecnologia through Programa Investigador FCT 2014 (IF/00576/2014) and Pluriannual funding to CCMAR (UID/Multi/04326/2013). This work is a contribution to the COST (European COoperation on Science and Technology) Action FA1301 "A network for improvement of cephalopod welfare and husbandry in research, aquaculture and fisheries (CephsInAction)" that partially funded the publication of this manuscript.

# ACKNOWLEDGMENTS

The authors thank Dr. Juan Fuentes, Dr. Pedro Guerreiro, Mr. João Reis and the students and staff at Ramalhete Station for valuable input and logistical assistance with the study.

#### REFERENCES

based on an initiative by CephRes, FELASA and the Boyd Group. Lab. Anim. 49, 1–90. doi: 10.1177/0023677215580006

**Conflict of Interest Statement:** The handling Editor declared a past coauthorship with one of the authors AS and states that the process nevertheless met the standards of a fair and objective review.

The other authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2017 Capaz, Tunnah, MacCormack, Lamarre, Sykes and Driedzic. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Corrigendum: Hypoxic Induced Decrease in Oxygen Consumption in Cuttlefish (Sepia officinalis) Is Associated with Minor Increases in Mantle Octopine but No Changes in Markers of Protein Turnover

#### Juan C. Capaz 1†, Louise Tunnah2†, Tyson J. MacCormack <sup>2</sup> , Simon G. Lamarre<sup>3</sup> , Antonio V. Sykes <sup>1</sup> \* and William R. Driedzic<sup>4</sup> \*

<sup>1</sup> Centro de Ciências do Mar do Algarve, Universidade do Algarve, Faro, Portugal, <sup>2</sup> Department of Chemistry and Biochemistry, Mount Allison University, Sackville, NB, Canada, <sup>3</sup> Département de Biologie, Université de Moncton, Moncton, NB, Canada, <sup>4</sup> Department of Ocean Sciences, Memorial University of Newfoundland, St. John's, NL, Canada

Keywords: European cuttlefish, Sepia officinalis, HSP70, octopine, polyubiquitinated protein, ventilation frequency

#### **A Corrigendum on**

#### **Hypoxic Induced Decrease in Oxygen Consumption in Cuttlefish (Sepia officinalis) Is Associated with Minor Increases in Mantle Octopine but No Changes in Markers of Protein Turnover**

by Capaz, J. C., Tunnah, L., MacCormack, T. J., Lamarre, S. G., Sykes, A. V., and Driedzic, W. R. (2017). Front. Physiol. 8:344. doi: 10.3389/fphys.2017.00344

In the original article, there was a mistake in **Figure 2B** as published. The right Y-axis of **Figure 2B**, concerning the amount of glycogen, was incorrectly printed resulting in glycogen levels 10-fold too low. The axis should read "0–8" and not "0–1." The corrected **Figure 2** appears below.

The authors apologize for this error and state that this does not change the scientific conclusions of the article in any way. The original article has been updated.

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2019 Capaz, Tunnah, MacCormack, Lamarre, Sykes and Driedzic. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

#### Edited and reviewed by:

Graziano Fiorito, Stazione Zoologica Anton Dohrn, Italy

#### \*Correspondence:

Antonio V. Sykes<EMAIL_ADDRESS>William R. Driedzic<EMAIL_ADDRESS>
†These authors have contributed equally to this work

#### Specialty section:

This article was submitted to Invertebrate Physiology, a section of the journal Frontiers in Physiology

Received: 11 December 2018 Accepted: 10 January 2019 Published: 30 January 2019

#### Citation:

Capaz JC, Tunnah L, MacCormack TJ, Lamarre SG, Sykes AV and Driedzic WR (2019) Corrigendum: Hypoxic Induced Decrease in Oxygen Consumption in Cuttlefish (Sepia officinalis) Is Associated with Minor Increases in Mantle Octopine but No Changes in Markers of Protein Turnover. Front. Physiol. 10:18. doi: 10.3389/fphys.2019.00018

oxygen saturation, 1 h) conditions. (A) octopine; (B) mantle free glucose and glycogen; (C) HSP70; (D) polyubiquitinated proteins. Statistical significance for octopine, HSP70, and polyubiquitinated proteins, was assessed with a 1-way ANOVA and for differences between glucose or glycogen levels with a t-test. N = 6 for all conditions except for free glucose in hypoxic mantle where N = 4. Differences between means or grouped means represent statistical difference (Tukey's multiple comparison test; P < 0.001). No differences were found in mantle free glucose and glycogen nor polyubiquitinated proteins (P > 0.05).

, Tanya J. Shaw<sup>3</sup>

,

# The Gastric Ganglion of *Octopus vulgaris*: Preliminary Characterization of Gene- and Putative Neurochemical-Complexity, and the Effect of *Aggregata octopiana* Digestive Tract Infection on Gene Expression

#### *Edited by:*

Elena Baldascino<sup>1</sup>

\* † , Giulia Di Cristina<sup>1</sup>

*Sylvia Anton, Institut National de la Recherche Agronomique (INRA), France*

#### *Reviewed by:*

*Sebastien Baratte, National Museum of Natural History, France Jean-Pierre Bellier, Shiga University of Medical Science, Japan*

#### *\*Correspondence:*

*Elena Baldascino<EMAIL_ADDRESS>These authors have contributed*

*†*

*equally to this work.*

#### *Specialty section:*

*This article was submitted to Invertebrate Physiology, a section of the journal Frontiers in Physiology*

*Received: 09 March 2017 Accepted: 20 November 2017 Published: 15 December 2017*

#### *Citation:*

*Baldascino E, Di Cristina G, Tedesco P, Hobbs C, Shaw TJ, Ponte G and Andrews PLR (2017) The Gastric Ganglion of Octopus vulgaris: Preliminary Characterization of Gene- and Putative Neurochemical-Complexity, and the Effect of Aggregata octopiana Digestive Tract Infection on Gene Expression. Front. Physiol. 8:1001. doi: 10.3389/fphys.2017.01001* Giovanna Ponte1, 4† and Paul L. R. Andrews 1, 4 *<sup>1</sup> Department of Biology and Evolution of Marine Organisms, Stazione Zoologica Anton Dohrn, Napoli, Italy, <sup>2</sup> Wolfson Centre*

, Perla Tedesco<sup>1</sup>

, Carl Hobbs <sup>2</sup>

*for Age-Related Diseases, King's College London, London, United Kingdom, <sup>3</sup> Centre for Inflammation Biology and Cancer Immunology, King's College London, London, United Kingdom, <sup>4</sup> Association for Cephalopod Research - CephRes, Napoli, Italy*

The gastric ganglion is the largest visceral ganglion in cephalopods. It is connected to the brain and is implicated in regulation of digestive tract functions. Here we have investigated the neurochemical complexity (through *in silico* gene expression analysis and immunohistochemistry) of the gastric ganglion in *Octopus vulgaris* and tested whether the expression of a selected number of genes was influenced by the magnitude of digestive tract parasitic infection by *Aggregata octopiana*. Novel evidence was obtained for putative peptide and non-peptide neurotransmitters in the gastric ganglion: cephalotocin, corticotrophin releasing factor, FMRFamide, gamma amino butyric acid, 5-hydroxytryptamine, molluscan insulin-related peptide 3, peptide PRQFV-amide, and tachykinin–related peptide. Receptors for cholecystokinin<sup>A</sup> and cholecystokininB, and orexin<sup>2</sup> were also identified in this context for the first time. We report evidence for acetylcholine, dopamine, noradrenaline, octopamine, small cardioactive peptide related peptide, and receptors for cephalotocin and octopressin, confirming previous publications. The effects of *Aggregata* observed here extend those previously described by showing effects on the gastric ganglion; in animals with a higher level of infection, genes implicated in inflammation (NFκB, fascin, serpinB10 and the toll-like 3 receptor) increased their relative expression, but TNF-α gene expression was lower as was expression of other genes implicated in oxidative stress (i.e., superoxide dismutase, peroxiredoxin 6, and glutathione peroxidase). Elevated *Aggregata* levels in the octopuses corresponded to an increase in the expression of the cholecystokinin<sup>A</sup> receptor and the small cardioactive peptide-related peptide. In contrast, we observed decreased relative expression of cephalotocin, dopamine β-hydroxylase, peptide PRQFV-amide, and tachykinin-related peptide genes. A discussion is provided on (i) potential roles of the various molecules in food intake regulation and digestive tract motility control and (ii) the difference in relative gene expression in the gastric ganglion in octopus with relatively high and low parasitic loads and the similarities to changes in the enteric innervation of mammals with digestive tract parasites. Our results provide additional data to the described neurochemical complexity of *O. vulgaris* gastric ganglion.

Keywords: digestive system, *Octopus vulgaris*, gastric ganglion, parasite, *Aggregata octopiana,* gene expression

#### INTRODUCTION

The gastric ganglion is a prominent feature of the peripheral nervous system in coleoid cephalopods. Gross morphological descriptions of the ganglion are available for Sepia officinalis (Alexandrowicz, 1928), Idiosepius paradoxus (Shigeno and Yamamoto, 2002), Octopus vulgaris (Young, 1967, 1971), and Eledone cirrhosa (Isgrove, 1909); illustration and a brief description during development is available for Sepioteuthis sepioidea (Shigeno et al., 2001) and Euprymna scolopes (Kerbl et al., 2013). In contrast to the single gastric ganglion in coleoid cephalopods, in Nautilus a pair of small ganglia distributing nerves to the viscera emerge from the visceral nerves (Owen, 1832).

The gastric ganglion (see original description for O. vulgaris in: Chéron, 1866; Bogoraze and Cazal, 1946) innervates most of the digestive tract, i.e., the crop, stomach, intestine, and caecum. It also connects with the central nervous system via the sympathetic nerves, the visceral nerves through rectal and intestinal nerves and through the abdominal nerves (Young, 1967). The complex structure of the gastric ganglion and its relationships support the view that it functions both independently and integrating information originating from, for example, the crop and intestine (Young, 1967), thus appearing to act not only as a simple relay but also as an integrative center (Andrews and Tansey, 1983). The intricate connectivity and complexity of the ganglion is further revealed by intense tubulinergic immunoreactivity of the neuropil (e.g., Shigeno and Yamamoto, 2002).

The well-defined innervation of the cephalopod digestive tract and the fact that it often hosts parasites (review in: Hochberg, 1983; Castellanos-Martínez and Gestal, 2013) raise the possibility that the presence of parasites may induce physiological responses (e.g., Gestal et al., 2002b) in the innervation, as occurs in mammals (see below).

In mammals, digestive tract pathogens (i.e., bacteria, viruses or parasites) can induce a range of responses including local inflammation, sensitization of visceral afferent nerves (peripheral terminal, cell body and central nervous system levels) and modulation of enteric nervous system (ENS) functionality (for review see: Halliez and Buret, 2015; Guarino et al., 2016; Obata and Pachnis, 2016). Examples are provided by the bacteria Campylobacter jejuni (Goehler et al., 2005), Clostridium difficile (Wadhwa et al., 2016) and Salmonella typhimurium (Gabanyi et al., 2016), rotavirus (Lundgren et al., 2000; Istrate et al., 2014) and the parasites Giardia duodenalis, Nippostrongylus brasiliensis, Trypanosoma cruzi and Trichinella spiralis (for review see Halliez and Buret, 2015). Mucosal damage of the digestive tract, such as occurs with an ulcer, can also produce sensory neuron sensitization (Bielefeldt et al., 2002), as can intestinal inflammation (Stewart et al., 2003).

With parasitic infections, changes observed in the gut innervation in rodents (mice or rats) include increased levels of the tachykinin substance P (e.g., Trichinella spiralis, Swain et al., 1992; Nippostrongylus brasiliensis, Masson et al., 1996), reduced acetylcholine release (e.g., Trichinella spiralis, Collins et al., 1989), and acute and chronic selective damage to the ENS (Trypanosoma cruzi, Campos et al., 2016). The sensitization of visceral afferent neurons and damage to the enteric innervation produced by pathogens contribute to post-infectious syndromes in humans (e.g., post-infectious gastroparesis in children, Naftali et al., 2007; post-infectious irritable bowel syndrome in adults, Schwille-Kiuntke et al., 2015; Wadhwa et al., 2016). Thus, the innervation of the mammalian digestive tract is affected both acutely and chronically by a range of pathogens including parasites.

In O. vulgaris the presence of parasites (review in: Hochberg, 1983; Castellanos-Martínez and Gestal, 2013) may induce responses either locally or systemically. The parasite most frequently found in octopus digestive tract is Aggregata octopiana (Estévez et al., 1996; Castellanos-Martínez and Gestal, 2013), a microscopic Coccidian, spore-forming, single-celled obligate intracellular parasite. It is one of the various species of Aggregata belonging to apicomplexan Protozoa (Apicomplexa: Aggregatidae). A. octopiana may reach incidences higher than 90% in some populations of O. vulgaris (e.g., West Mediterranean, Mayo-Hernández et al., 2013). In the digestive tract of octopus, A. octopiana is found in both non-cuticularized (caecum and intestine), and cuticularized (esophagus and crop) structures, in the digestive gland and other nearby organs (Gestal et al., 2002a,b). In infected animals, cysts are visible with the naked eye as small white patches embedded in the muscular wall of the digestive tract (Mayo-Hernández et al., 2013). Therefore, they are in close proximity to the enteric neurons located in the serosa and between the circular and longitudinal muscle layers of the digestive tract in cephalopods (Alexandrowicz, 1928; Graziadei, 1960).

Histological and ultrastructural lesions associated with A. octopiana infection include hypertrophy with nuclear displacement of the host cell, inflammation, phagocytosis, ulceration and final damage of the organ architecture (Gestal et al., 2002a). Aggregata appears to be particularly abundant in the caecum and intestine in the octopus, and has been proposed to be responsible for a "malabsorption syndrome" (Gestal et al., 2002b). A. octopiana has also been suggested to induce a systemic immune response (Castellanos-Martínez et al., 2014a; Gestal and Castellanos-Martínez, 2015).

# AIM OF THIS STUDY

The effects of the lesions associated with A. octopiana infection in O. vulgaris have been explored at different levels, but the impact on neural structures has never been investigated. Here we ask whether Aggegata affects one component of the digestive tract innervation, namely the gastric ganglion. Our hypothesis is based on the assumption that the gastric ganglion is connected, through both sensory and motor neurons, to all regions of the digestive tract, which can be affected by Aggregata, and therefore the damage and local inflammation induced by the parasite will cause functional modification reflected in gene expression. The neurons of the gastric ganglion could also be affected by substances (e.g., TNFα) released as components of the systemic immune response to Aggregata (Castellanos-Martínez et al., 2014a,b).

To explore our working hypothesis and to expand current knowledge of the neural complexity of this ganglion, we: (i) surveyed the neurochemical diversity of the gastric ganglion in O. vulgaris by a combination of approaches including in silico molecular characterization and immunohistochemistry with a particular focus on ligands and receptors likely to be involved in neurotransmission; (ii) used real time RT-qPCR to compare the expression of selected target genes in the gastric ganglion from octopuses with relatively "high" and "low" Aggregata parasite loads, to provide insights into the potential impact of infection on the control of the digestive tract.

#### MATERIALS AND METHODS

#### Animals and Tissue Sampling

Octopus vulgaris Cuvier, 1797 (Mollusca, Cephalopoda) of both sexes (males, N = 10 − body weight: mean ± SEM = 185 ± 22 g; females, N = 12 − body weight: mean ± SEM = 272 ± 19 g) were obtained from local fishermen (Bay of Naples, Italy). Twelve octopuses were used for immunohistochemistry, and an additional 10 utilized for analysis of the potential effects of Aggregata infection on the gastric ganglion (see below). All animals originated from a larger sampling study measuring the incidence of octopus' parasite load (A. octopiana) in the Mediterranean.

Killing animals solely for tissue removal does not require authorization from the National Competent Authority under Directive 2010/63/EU and its transposition into national legislation. Samples were taken from local fishermen, by applying humane killing following principles detailed in Annex IV of Directive 2010/63/EU as described in Fiorito et al. (2015). In brief, octopuses were immersed in freshly made 3.5% magnesium chloride hexahydrate (Sigma Aldrich, CAS Number: 7791-18-6) dissolved in sea water**.** After 30 min immersion, the animals were unresponsive to handling and a noxious mechanical stimulus, and ventilation completely stopped; killing was completed by destruction of the brain.

#### Tissue Removal

All dissections were carried out on an ice bed. The digestive tract was approached via incision of the dorsal mantle and of the capsule covering the digestive gland. The gastric ganglion was identified on the right-hand side at the junction of the crop, stomach, caecum and intestine (**Figure 1**). The ganglion was removed by transection of nerve trunks projecting to adjacent structures and cutting the connective tissue capsule adherent to the serosa of the digestive tract. The ganglion was placed initially in either fixative (for immunohistochemistry) or RNA Later Stabilization Solution (RNALater, Thermo Fisher Scientific; for gene expression studies) as indicated below.

In addition, the entire digestive tract was removed by transection of the esophagus as it exited the brain and the rectum at the level of the anal sphincter. Blunt dissection freed the gastric ganglion from adherent structures.

For all animals, the digestive tract was inspected for the presence of white cysts indicative of A. octopiana infection and rapidly frozen on dry ice and then stored at −20◦C for subsequent quantification of the magnitude of A. octopiana infection (see below).

# Immunohistochemistry and Staining

Excised gastric ganglia from animals with no visible Aggregata infection were immersed in fixative appropriate for each of the employed antisera and/or treated for the specific purpose of the required staining (**Table 1**). In most cases for paraffin and cryostat sections ganglia were placed in 4% paraformaldehyde (PFA) in seawater (4◦C for 2 h). After fixation, the tissue was treated as briefly summarized below.

For fluorescence immunohistochemistry (IHC) on paraffin tissue, sections were deparaffinized, rehydrated and subjected to heat-mediated antigen retrieval in Sodium Citrate buffer (pH

FIGURE 1 | The gastric ganglion of *Octopus vulgaris*. (Left panel) A dissected digestive tract from an octopus. From top (anterior) to bottom (posterior): Posterior salivary glands (Psg), crop (Cr), esophagus (Oes), stomach (St), caecum (Cae), and intestine (Int). Within the square (see magnification in the Right panel), the arrow points to the gastric ganglion. (Right panel) In higher magnification, the same structures are identified together with the gastric ganglion (GG) and surrounding nerve bundles (see black lines to highlight their relative positions) running toward the stomach and crop (up), and toward the caecum and digestive duct (bottom left) or toward the intestine (bottom right). The proximal part of the intestine (Pr. Int) and the cut end of the digestive duct (DD) are also evident. Scale bars: 500µm.

#### TABLE 1 | List of antibodies utilized in this study and their source.

*Name and abbreviation and type of antibody (monoclonal: M, polyclonal: P) utilized are included together with supplier and catalog number. Cited references refer to previous studies where the same antibody (or with identical epitope) has been utilized for nervous tissue in cephalopods.*

*<sup>a</sup>Antibody applied after antigen retrieval as described by Herculano-Houzel and Lent (2005): 1 h (70*◦*C) in 0.2 M boric acid solution (pH 9). The sections were then washed in PBS and incubated with a primary antibody against NeuN (rabbit polyclonal; Cy3 conjugate).*

*<sup>b</sup>To detect cChAT-ir we followed recommendations provided by Casini et al. (2012).*

6.0). Sections were blocked in 10% goat serum (2h at RT) and then incubated at 4◦C with primary antibody overnight. After washing (PBS, 3 × 5 min) tissues were incubated with AlexaFluor-conjugated secondary antibodies as appropriate for 2 h at 37◦C in the dark. For the final 15 min, DAPI was added for detection of DNA/nuclei. Tissue was again washed in PBS (3 × 5 min), and then mounted with fluorescent mounting medium (FluoromountTM Sigma Aldrich).

For NeuN detection we applied antigen retrieval to paraffin sections as described by Herculano-Houzel and Lent (2005), i.e., boric acid solution (see **Table 1** for details) and then incubated with a primary antibody against NeuN (rabbit polyclonal; Cy3 conjugate).

To detect cChAT immunoreactivity we followed the standard protocol utilized for octopus (Casini et al., 2012).

The omission of each primary antibody was used as a negative control for each immunostaining procedure.

Classic histological staining was utilized to examine the general organization of the octopus' gastric ganglion. We applied Hematoxylin and Eosin (H&E) on deparaffinized/rehydrated tissues incubated in Mayer's hematoxylin (Sigma) for 5 min, then washed in running tap water (5 min), followed by incubation with Eosin Y (Sigma) solution (1 min), and then rapidly dehydrated, cleared in xylene, and mounted (Mayer, 1893; e.g., Shigeno et al., 2001). Serial sections (50µm) were also collected on chromealum-gelatin-coated slides, and stained using with Picro-Ponceau with hematoxylin following Kier (1992).

Classical histological sections were examined using brightfield microscopy (Zeiss Axiocam 2, with Zen-Blue software) and photographed with a Zeiss105 color camera. For IHC we utilized a Leica DMI6000 B inverted microscope and a Leica TCS SP8 X confocal microscope (Leica Microsystems, Germany). Tile Zstacks were performed using a 0.2µm step size. Images were processed using LAS X software (Leica Microsystems, Germany). IHC figures have been assembled following guidelines for color blindness provided by Wong (2011).

#### Assessment of *Aggregata octopiana* Infection

The entire digestive tract (esophagus to rectum) of O. vulgaris (N = 10) was removed immediately post mortem, weighed and after vigorous washing in homogenization buffer processed according to Gestal and co-workers (Gestal et al., 1999; see also Gestal et al., 2007; Tedesco et al., 2017). In brief, after homogenization, sporocysts of A. octopiana were counted by a Neubauer chamber and their number expressed as sporocysts per gram of tissue. Samples considered in this study were taken from a larger sampling study aimed to describe possible differential parasite loads in different octopuses. We selected samples to assure a significant difference in parasite load between animals (see section Results).

#### Transcriptome Analysis and *in Silico* Characterization of the Gastric Ganglion

A preliminary characterization of the complexity of the gastric ganglion of O. vulgaris was conducted utilizing recent transcriptome data derived from Drs. R. Sanges' and G. Fiorito's Research Groups at the Stazione Zoologica Anton Dohrn (Napoli, Italy). We utilized a dataset based on two separate RNA-seq studies (Petrosino, 2015; Zarrella et al., unpublished data) carried out on O. vulgaris central nervous system (i.e., optic lobes, supra-esophageal and sub-esophageal masses), distal extremities of arm (including muscular and nervous tissues), stellate and gastric ganglia (for technical details see: Musacchia et al., 2015; Petrosino, 2015). The resulting transcriptome (Drs. R. Sanges and G. Fiorito Labs: Stazione Zoologica Anton Dohrn; see also Petrosino, 2015) identified more than a hundred thousand transcripts from different neural structures, significantly extending previously available transcriptome data for this species (Zhang et al., 2012).

We applied a biased strategy, based on Gene Ontology (http:// www.geneontology.org/), to these datasets and mined for genetranscripts potentially involved and/or implicated in the response to infection, inflammation, immune/stress responses.

#### Analysis of Gene Expression RNA Extraction and cDNA Synthesis

Gastric ganglia were dissected from animals and stored in RNA Later Stabilization solution for 24 h (+4 ◦C) and transferred to −80◦C until further processing. Total RNA was extracted using SV Total RNA Isolation System (Promega, Z3100) according to manufacturer instructions. Quality and quantity of extracted RNA was assessed through UV absorption measurements (Nanodrop ND-1000 UV-Vis spectrophotometer, Nanodrop Technologies). Absence of DNA contamination was verified through PCR (ubiquitin primers) followed by gel electrophoresis. For cDNA synthesis, 500 ng of total RNA from each sample were processed with iScript-cDNA Synthesis Kit (Bio-Rad, 1708891).

#### Primer Design: Efficiency and Specificity

Primers were designed by Primer3Plus software (www. bioinformatics.nl/primer3plus) using target sequences deduced from O. vulgaris transcriptome (Petrosino, 2015). Primer parameters were set to 20 nucleotides in length, product size 100–200 base pairs and melting point 58–60◦C. Primers were also analyzed with the Multiple Primer Analyzer<sup>1</sup> to estimate presence of, and possibly avoid, primer-dimers.

The efficiency of each pair of primers (**Table 2**) was calculated according to standard methods curves, and following Sirakov et al. (2009). Briefly, five serial dilutions (1:5, 1:10, 1:20, 1:40, 1:80) of a standard sample were made to determine the efficiency of reactions conducted with each pair of primers. Standard curves were generated for each sample/gene combination using the Ct value vs. the logarithm of each dilution factor (Pfaffl et al., 2002; Radonic et al., ´ 2004).

Each amplification reaction was conducted in a volume of 25 µl containing: 2 µl of diluted cDNA template, 2.5 µl of 10× PCR reaction buffer (Roche), 2.5µl of dNTP mix (0.2 mM), 1µl of each primer (25 ρmol/µl), 0.25µl of Taq DNA polymerase (5U/µl), and sterile H2O. The amplification cycles were conducted by Peltier Thermal Cycler PTC-200 (MJ Research). After denaturation at 95◦C (2 min) 34 amplification cycles were carried out as follows: denaturation (94◦C, 15 s), annealing (60◦C, 30 s), extension (72◦C, 1 min). Finally, an extension cycle was carried out at 72◦C for 7 min to complete

<sup>1</sup>https://www.thermofisher.com/it/en/home/brands/thermo-scientific/molecularbiology/molecular-biology-learning-center/molecular-biology-resource-library/ thermo-scientific-web-tools/multiple-primer-analyzer.html

#### TABLE 2 | *O. vulgaris* genes (*n* = 33) identified and utilized in this study.

*(Continued)*

*Transcripts are given together with valid protein and gene names, and Uniprot accession number (AN); GeneBank accession numbers for O. vulgaris orthologs of these genes are indicated in parenthesis. Whenever available and for nucleotidic sequences that have been validated we provide GenBank accession number (AN, in parenthesis). The table also include sequences of primers (n* = *24, Forward and Reverse) used for RT-qPCR experiments, amplicon size (AS) and efficiency (E). Gene ontology (GO) annotation for biological processes, functions and components are included. GO identifiers marked with* \**refer to cases where GO were attributed following the identification of similar proteins known in vertebrates or invertebrates to have analogous functions and when GO does not provide any result based on the valid Protein Name.*

all the strands. PCR products were run on 2% agarose gel in TBE buffer 0.5× (45 mM Tris-borate, 1mM EDTA) and detected expected bands were isolated. DNA was extracted using GenElute Gel Extraction Kit (Sigma-Aldrich, NA1111) and analyzed using an Automated Capillary Electrophoresis Sequencer 3730 DNA Analyzer (Applied Biosystems).

#### Real-Time qPCR

In order to analyze expression levels of specific genes of interest, a panel of putative reference genes was first screened to find the most stable genes for these experimental conditions (see the approach utilized for octopus in Sirakov et al., 2009). In our experiments, we utilized: eukaryotic translation initiation factor 4 (EIF4G1), LIM and SH3 domain protein (F42H10.3), lamin-B1 (Lmnb1), cytoplasmic FMR1 (Sra-1), ubiquitin-40S ribosomal protein S27a (RPS27A), elongation factor 1-alpha (eef1a), 40S ribosomal protein S18 (RPS18). The gene expression stability of the candidate reference genes for our samples was evaluated with BestKeeper (Pfaffl et al., 2002) and NormFinder (Andersen et al., 2004), following Sirakov et al. (2009). We identified the three most stable reference genes as Lmnb1, Sra-1 and RPS27A.

For gene expression experiments, samples from 10 octopuses were processed in triplicate. Polymerase chain reactions were carried out in an optical 384-wells plate with Applied Biosystems ViiA7 (Life Technologies) using Fast SYBR Green Master mix (ThermoFisher Scientific) to monitor dsDNA synthesis. Reactions (total volume: 10 µl) contained: 1 µl cDNA, 5 µl SYBR Green Master mix reagent, 4 µl of forward and reverse primers mix (0.7 pmol/µl each). The following thermal profile was used: 95◦C for 10 min; 95◦C for 15 s, 60◦C for 1 min, 40 cycles for amplification; 72◦C for 5 min; one cycle for melting curve analysis, from 60◦ to 95◦C to verify the presence of a single product. Specificity of PCR products was checked by melting curve analysis followed by gel electrophoresis and DNA sequencing. PCR data were analyzed using the ViiATM 7 Software (Life Technologies) to determine cycle threshold (Ct) values. Each assay included a no-template control for every primer pair. All sequences have been deposited in GenBank after validation (**Table 2**).

#### Data Analysis

Relative expression of the genes of interest, identified by in silico analysis of the octopus transcriptome, in the gastric ganglion and other tissues was analyzed through hierarchical clustering and principal component analysis (PCA) using Multi-experiment Viewer (MeV) software (Saeed et al., 2003). Quantitative real-time PCR experiments, carried out to evaluate the response of the octopus gastric ganglion to different levels of A. octopiana infection ("high" vs. "low" parasite load), were analyzed through Multivariate Analysis of Variance (MANOVA) following Tsai and Chen (2009). Gene expression changes are expressed as Log<sup>2</sup> fold-changes following common practice (Friedman et al., 2006; Fundel et al., 2008). For all statistical analyses we used SPSS (rel. 18.0, SPSS Inc. - Chicago, 2009), with the exceptions mentioned above, and following Zar (1999). All tests were two-tailed and the alpha was set at 0.05.

#### RESULTS

#### *In Silico* Comparison of Gastric Ganglion Transcripts with Other Tissues

**Figure 2** illustrates the transcriptional profiles of 33 genes expressed in O. vulgaris gastric ganglion (**Table 2**), selected through Gene Ontology (GO) Biological Functions search of the available transcriptome data. In brief, we counted over 65,000 nucleotide transcripts selectively expressed in the octopus' gastric ganglion. Our biased GO searching strategy probed for transcripts considered of particular relevance to the response to infection (e.g., neurotransmitter-synthesizing enzymes, receptors for transmitters and hormones), or implicated in inflammatory and/or immune/stress responses (e.g., superoxide dismutase, tolllike receptors, fatty acid-binding homolog 9); this provided a reduced number of potential candidate genes.

A more comprehensive analysis of the transcriptome fingerprints of O. vulgaris gastric ganglion is beyond the scope of this study, and this list should be considered preliminary.

We found that in silico relative abundance (counts) of the 33 transcripts in the octopus gastric ganglion varied. These ranged from <1 CPM (endothelin converting enzyme 1) to >1,000 CPM for fatty acid-binding homolog 9 (Lbp-9), the highest count for any transcript considered in any tissue. We

FIGURE 2 | Heatmaps and hierarchical clustering of gene expression levels in octopus brain (OL, SEM, SUB), stellate (SG) and gastric (GG) ganglia, and arm tip (TIP). Thirty-three (Left panel) and 24 transcripts (Right panel) mined from *Octopus vulgaris* RNAseq data were selected with a focus on neurotransmitters, signaling machinery and inflammation. Data were normalized per gene and subsequently analyzed by hierarchical clustering of the samples (tissues) and transcripts. Differences in the relative expression and clustering revealed a differential pattern between the gastric ganglion and other tissues considered independently from the number of transcripts included in this preliminary *in silico* characterization of octopus gastric ganglion. See Table 2 and text for details.

also found 16 gene transcripts with counts <10 CPM, and nine in the range 10–100 CPM (allograft inflammatory factor, SCPRPamide, Protein PRQFV-amide, glutathione S-transferase, glutathione peroxidase, superoxide dismutase, peroxiredoxin 6 protein, glutathione synthase-like isoformX3; hypothetical protein OCBIM). Finally, 6 of the 33 genes considered had transcript counts in the range 100–1,000 CPM (dopamine betahydroxylase, tachykinin related peptide, Rab effector Noc2, cephalotocin, molluscan insulin-related peptide 3, protein singed). In silico relative expression of these 33 transcripts are overviewed in **Figure 2**.

A subset of 24 of the 33 genes (**Table 2**) was validated through real time RT-qPCR (see also **Figure 6**). In addition, and to confirm homogeneity between the two sub-sets of O. vulgaris gastric ganglion transcripts, we produced heatmaps and performed hierarchical clustering analysis (**Figure 2**). Differences were identified in the transcriptional fingerprint of O. vulgaris gastric ganglion when compared with other tissues, namely the stellate ganglion, arm tip and brain regions (**Figure 2**). The differences emerged independently from the number of transcripts considered (i.e., either 33 or 24 genes; **Figure 2**).

Principal Component Analysis (PCA) confirmed that the gastric ganglion profile differs from the other tissues (**Figure 3**). The two first principal components accounted for less than 50% of the total variance, and the cut-off eigenvalue (set to 1) was achieved only with principal component 5 (for PC analysis of either 33 or 24 transcripts, data not shown). Nevertheless, the gastric ganglion segregated into a different quadrant from the stellate ganglion. Furthermore, the analysis showed that the arm tip and the regions belonging to octopus' central nervous system also segregated in different quadrants (**Figure 3**), thus confirming that the genes considered here have a specific expression profile, and revealing that the three different brain regions (optic lobes and supra- and sub-esophageal masses) cluster together, but distant from the peripheral tissues considered.

### Extending Current Knowledge of *O. vulgaris* Gastric Ganglion Structural Complexity

O. vulgaris gastric ganglion is an ovoid structure prominently located at the junction of the crop, stomach, caecum and proximal intestine at the point where they share a common lumen (**Figure 1**). The well-circumscribed nature of the ganglion was particularly evident following histological examination, which confirmed previous data reporting that it is almost entirely encapsulated in connective tissue (Bogoraze and Cazal, 1946; **Figures 4A–C**). The ganglion itself consists of very densely packed nerve cell bodies distributed in a cortical layer of the oblate spheroid structure, with axons coursing centrally, bundling, and allowing the ganglion to be connected with other structures including the subjacent digestive tract (**Figures 4A–C**). Following Bogoraze and Cazal (1946), the peripheral layer of cells appears relatively larger than those that are more closely distributed toward the internal neuropil (see also Young, 1971).

Numerous cells also appear in the neuropil. A group of them are clustered together, toward the center of the ganglion, creating

FIGURE 3 | Principal Component Analysis of gene expression data based on transcript levels (*in silico*, see Figure 2) of 33 genes selected for this study (see Table 2 for full list). Data are plotted for tissue taken from three (*N* = 3) different animals. Dissimilarity between peripheral and central nervous system tissues emerged from the analysis with a clear distinction of the octopus gastric ganglion signature from other peripheral structures (i.e., stellate ganglion and arm). See text for details.

a wall apparently separating it into two sections (**Figure 4J**, arrowheads; see description in Young, 1971). In addition, some cells are dispersed in the neuropil and have been described by Bogoraze and Cazal (1946) as forming a glio-vascular network (**Figures 4E,I**). In our samples, we found cells with somata diameters ranging from 7 to 30µm.

In the following section, a further description of the cellular components and fibers constituting the octopus gastric ganglion, together with information derived from attempts to localize modulators and other characteristic markers, is provided through immunohistochemistry (IHC). The results should be considered preliminary. This because the currently available commercial antibodies we utilized are not designed for octopus or cephalopods. However, as shown in **Table 1** (consider also exceptions therein) these have been already applied to cephalopod tissues in a series of studies, despite that in some cases standardized ways to biologically validate them have not being applied. Therefore, observations provided below should be considered as an indication of "-like immunoreactivity" (referred hereunder as like-IR or IR). These limitations are considered further in the Discussion.

We utilized NeuN for the first time in octopus. It positively identified the great majority of cells in the gastric ganglion (**Figures 4J–M**) confirming its general architecture (see above). In addition, there were a few cells negative for the NeuN-antibody (**Figure 4M**, arrowheads), suggesting also the existence of supporting cells (e.g., glia-like cells) in the gastric ganglion. A complex organization of fibers was revealed through neurofilament-IR; these appeared mostly ordered and compacted into bundles in the part proximal to the sympathetic (**Figure 4D**, sn) nerve (i.e., superior and dorsal bundles, sensu Bogoraze and Cazal, 1946). On the other hand, fibers appeared greatly intertwined toward the posterior area of the neuropil

the ganglion, with its cortical layer of neural cells (Cell layer) and internal neuropil. Nerve bundles are highlighted; in (A) intestinal (in) and sympathetic (sn) nerves are indicated; staining also revealed blood vessels (bv). Connective tissue and the cell layer surrounding and enveloping the ganglion is clearly identified after Picro-Ponceau with hematoxylin staining (C). (D–M) Fluorescent immunohistochemistry for (in green) neurofilament (D–F), acetylated alpha-Tubulin (G–I), and NeuN (J–M), DAPI is utilized as nuclear stain (magenta). Neurofilament marks the inner neuropil (D) and the intricate organization of fibers (enlargement in E), with some neural processes entering the cellular layer (enlargement in F). Acetylated alpha-tubulin (G–I) shows the external group (external layer) of cells larger than those distributed internally (internal layer) (H) with several processes progressing toward the internal neuropil (H,I). NeuN and nuclear stain (DAPI) merged image (J) are provided as separate channels in (K,L), respectively for areas corresponding to the white rectangle. A cell wall dividing the neuropil of the ganglion is clearly visible (J, arrowheads). Higher magnification to show the neurons positive for NeuN (green), DAPI (magenta), and merged channels in (M). The cells not positive for NeuN are indicated with arrowheads. Scale bars: 500µm (D,G,J,K,L); 200µm (A–C); 100µm (E,F,H,I); 10µm (M).

(**Figure 4D**, left rectangle in; enlargement in **Figure 4E**), before the emergence of the ventral nerve bundles (see also intestinal nerves in Young, 1967). It is in this area that neurofilament-IR revealed fibers forming polygonal, round or ovoid networks resulting from the convergence of numerous fiber bundles (**Figures 4D–F**).

Using acetylated alpha-tubulin antibody we found positive cells and fibers (**Figures 4G–I**). The fibers appeared intertwined and arranged in different directions and orientations. Most of them appeared to emerge from larger cells forming networks where cells appeared dispersed into bundles of different size and complexity (**Figures 4F,H**).

To gain a broader understanding of the molecular complexity and signaling potential of the gastric ganglion, the expression of a range of putative neurotransmitters was also analyzed by IHC.

We observed numerous nerve fibers positive for tyrosine hydroxylase antibody (**Figures 5A,B**). These are identified as ordered bundles in the neuropil of the ganglion and toward the anterior pole in areas belonging to the sympathetic nerve bundles. This pattern contrasts with the diffuse and apparently disorganized arrangement observed toward the posterior end, corresponding to the area where neurofilament-like IR revealed intricate round (or ovoid) network of fibers.

A diffuse, but evident and widely distributed IR-signal occurred in sections using octopamine antibody. Octopamine immunoreactivity revealed small "button-like"-vesicles (**Figures 5C,D**) widely distributed in the cytoplasm and neuronal processes, again suggesting the existence of an intricate network.

5-hydroxytryptamine-like positive nerve fibers (**Figures 5E,F**) were clearly visible in the neuropil, in some cases forming a clustered network (**Figure 5F**, arrowheads). Furthermore, we identified several processes originating from larger neurons and progressing toward the neuropil, in some cases surrounding cells belonging to the internal layer of the ganglion (**Figure 5F**, arrows).

Common type of choline acetyltransferase (cChAT) antibody identified a few sparse, but clustered positive cells (**Figures 5G,H**), and several positive fibers dispersed in the neuropil.

GABA-like IR revealed an intricate widely distributed reticulum surrounding the great majority (almost all) of cells; this appeared particularly evident in the cells belonging to the more external layers of the cortical zone (**Figure 5I**). It overlapped with the noradrenaline-like IR we observed (**Figure 5J**), although appearing less intricate when compared with GABA-like IR. Noradrenaline-IR is also found in fibers of the nerve bundles (**Figure 5K**).

Corticotrophin releasing factor (CRF-like IR) was identified in a subset of the cells within the cortical layer of the gastric ganglion (**Figure 5L**) and also in a few CRF-like IR positive nerve fibers.

FMRFamide-like immunoreactivity was widely distributed in the ganglion with numerous positive fibers identified in various parts of the neuropil and in several cells distributed in the surrounding cortical layer (**Figures 5M,N**). In several areas, strong FMRFamide-like IR was seen in the neuropil in a cluster of fibers forming a beaded appearance (**Figure 5M**). Furthermore, positive FMRFamide-like fibers have been identified in the nerves that connect the gastric ganglion with other structures.

Various cells of the internal zone of the cortical layer of the gastric ganglion were positive for tachykinin–like immunoreactivity (**Figure 5O**). We also observed numerous tachykinin-like positive fibers both in the neuropil and nerves (**Figure 5P**).

#### Gene Expression in the Gastric Ganglion as a Result of *Aggregata octopiana* Infection

We found diversified expression of the selected 24 genes in the gastric ganglion after RT-qPCR experiments linked to parasite load in the digestive tract samples (**Figure 6**; see also **Table 3**). We compared gene expression in gastric ganglia of octopuses with an elevated number of A. octopiana sporocysts ["high," sporocysts/tissue(g): median = 1.37<sup>∗</sup> 10<sup>6</sup> , 95% CI ± 1.80; N = 5] in their digestive tract, with those from O. vulgaris with a relatively low parasite load ["low," sporocysts/tissue(g): median = 0.26<sup>∗</sup> 10<sup>6</sup> , 95% CI ± 0.21; N = 5; parasite incidence, "high" vs. "low": Z = 2.6, N1, N2 = 5, p = 0.009 after Mann-Whitney U test]. In the latter group, we counted fewer than 6,000 sporocysts per gram of tissue; in "high" group we found the highest value of Aggregata sporocysts [sporocysts/tissue(g) = 5.8<sup>∗</sup> 10<sup>6</sup> ]. In three out of ten animals we found a very few (<3/octopus) larval forms of other parasites, i.e., cestodes and nematodes. There is no evidence for effects of these parasites on the functioning of the cephalopod digestive system, to the best of our knowledge (Hochberg, 1990).

We observed a significant change in gene expression in the gastric ganglia in response to infection [high vs. low-−24 genes, MANOVA: F(4, 7) = 202.59, p < 0.001]. **Figure 6** plots gene expression as fold changes observed in octopus gastric ganglia from "high" relative to those sampled from "low" parasite load animals. Seventeen out 24 genes had significant pairwise changes (after Bonferroni post-hoc comparison; **Figure 6**; **Table 3**). Under these computational circumstances, six genes increased their relative expression (high vs. low: Sn, Nfκb2, Cckar, SCPRP, Serpinb10, Tlr3) while decreased gene expression was observed in the others (n = 11; high vs. low: Litaf, Dbh, OPR, CT, Mirp, PRQFV, Tk, Sod1, Prdx6, Gpx1, Rph3al). Furthermore, genes for Ov-Nuclear factor NF-κB p100 subunit and Ov-Toll like receptor 3 were found with at about one-fold increase (**Figure 6**). At least a one-fold decrease was observed for Ov-Lipopolysaccharideinduced tumor necrosis factor-alpha factor, Ov-Molluscan insulin-related peptide 3, Ov-Superoxide dismutase [Cu-Zn] (Ov-Sod1), and Ov-Rab effector Noc2 (**Figure 6**).

#### DISCUSSION

This study provides an update of the knowledge on the complexity of the gastric ganglion of the cephalopod mollusc O. vulgaris using a combination of immunohistochemical and gene expression analyses. Recent transcriptome analyses of octopus and other cephalopods (e.g., Zhang et al., 2012; Albertin et al., 2015; Petrosino, 2015; Salazar et al., 2015; Liu et al., 2016; Liscovitch-Brauer et al., 2017; Tian et al., 2017) have greatly facilitated this study. The data we accessed (R. Sanges and G. Fiorito Laboratories O. vulgaris transcriptome; see Materials and Methods) enabled characterization of the molecular fingerprint of octopus' gastric ganglion. Despite limitations imposed by the set of transcripts considered (33 out over 60,000 in total),

FIGURE 5 | Neurochemical complexity of *Octopus vulgaris* gastric ganglion. Fluorescent immunoreactivity (in green) for: Tyrosine Hydroxylase (TH, A,B), Octopamine (OA, C,D), 5-hydroxytryptamine (5-HT, E,F), common Choline Acetyltransferase (cChAT, G, H), GABA (I), Noradrenaline (NA, J, K), Corticotrophin releasing factor (CRF, L), FMRF-amide (M,N), and Tachykinin (TK, O,P); DAPI is utilized as nuclear stain (magenta, except in N–P). Co-localization with acetylated alpha-tubulin (AcTub) is shown in (A,E,N,O,P) with neurofilament (NF) in (C,G). TH-IR fibers as they appear in the internal neuropil of the gastric ganglion (A) co-localized with AcTub (gray). The white square area in (A) is enlarged in (B) to show the strong TH immunoreactivity at the level of neural processes. (C) The intricate network of octopamine positive fibers and button-like positivity as seen in the neuropil and cellular layer (colocalized with NF, gray). The high magnification in (D) shows details of octopamine-IR fibers inside the cellular layer and their prolongation within the internal neuropil. (E) 5-HT-IR and AcTub (gray) revealed fibers in the internal neuropil with several processes progressing toward the cellular layer. The high magnification detail in (F) shows some of these processes in the cellular layer (arrows), and some clustered network fibers (arrowheads). (G) The common Choline Acetyltransferase immunoreactivity seen in cells and fibers, co-localized with NF (gray). The magnification in (H) to highlight cChAT-IR in cells. GABA-IR (I) identified an intricate network surrounding the cells of the cortical layer; a magnification is provided in the enlargement. (J) Noradrenaline immunoreactivity revealing an intricate reticulum surrounding cells of cellular layer, and at the level of the sympathetic nerve (K). CRF-IR positive cells (L) and detail in the higher magnification (square) of the cellular layer. FMRFamide-IR fibers are observed in the intestinal nerve (transverse section, M). FMRFamide-IR fibers in the neuropil of the gastric ganglion (N) with processes and cells in the cellular layer (co-localized with AcTub, magenta). Tachykinin-like IR in cells and fibers in the neuropil (O,P); co-localization with AcTub (magenta). See text for details. Scale bars: 100µm (A,B,D,E,H,I,J,K,L); 50µm (C,G,F,M,N,O,P); 25µm (higher magnification in L,I).

in silico gene expression analysis revealed that the ganglion is clearly distinguishable from other central and peripheral nervous system tissues, when the same set of genes is considered. This is confirmed by PCA (**Figure 3**), with the gastric ganglion segregated into a different quadrant in respect to the stellate ganglion, and the arm tip and the octopus' brain structures clustered in other quadrants (**Figure 3**).

Below we will first summarize our data and the literature to depict the neurochemical complexity of O. vulgaris gastric ganglion, and then discuss the effects of Aggregata infection on gene expression in the ganglion.

# *O. vulgaris* Gastric Ganglion: A Contribution to the Understanding of Its Neural Complexity

In O. vulgaris the gastric ganglion appears as a white, ovalshaped, encapsulated structure (see **Figure 1**) about 3 mm long (in a 500 g body weight animal Andrews and Tansey, 1983). It is located on the serosa of the digestive tract at the junction of the crop, stomach, caecum and proximal intestine (Bogoraze and Cazal, 1946; Young, 1967). The cephalopod gastric ganglion is characterized by a dense neuropil surrounded by cell bodies, encapsulated by connective tissue. This structure was apparent from the H&E staining (**Figure 4C**), which also showed axon bundles exiting the ganglion for adjacent regions of the digestive tract, and confirmed previous descriptions (see Bogoraze and Cazal, 1946; Young, 1967, 1971). The dense layer of nerve cell bodies surrounds a central neuropil, with axons forming the nerve bundles linking, through an intricate network, to the adjacent structures: crop, stomach, caecum, hepatic ducts and intestine—(Bogoraze and Cazal, 1946; Young, 1967). The major nerve bundles have been identified as: superior bundle (3 nerves: anterior- and posterior-gastric, and gastro-esophageal nerves), dorsal bundle (spreading in a fan innervating the caecum), and ventral bundle (3 nerves) emerging from the posterior end (**Figure 4G**: in) of the gastric ganglion and projecting toward the intestine (see descriptions in Bogoraze and Cazal, 1946; Young, 1967).

We observed nerve cell bodies larger (external layer) than those closer (internal layer) to the neuropil (**Figures 4H**, **5N**), in analogy to other ganglia in octopus as described by Bogoraze and Cazal (1944; 1946; see also Young, 1971). However, very small cells (less than 5µm diameter) were not confirmed by our observations.

In the neuropil a dividing partition made of cell bodies (**Figure 4J**, arrowheads) was also noted, as reported by Young (1967, 1971). In Nautilus there are paired gastric ganglia (Owen, 1832) and we hypothesize that the partition observed in octopus' gastric ganglion is a remnant of the fusion of paired ganglia in an ancestral cephalopod to form a single ganglion in modern cephalopods.

*Results after MANOVA are computed after Bonferroni correction. See Figure 6 for direction of changes.*

*Bolded p values show statistically significant differences.*

The characterization of O. vulgaris gastric ganglion was further extended by identifying neuronal markers that can be utilized for studying the morphology of the cephalopod ganglia moving forward.

We identified cell bodies using NeuN antibody, a neuronal nuclear marker known to recognize neurons in the both the central and peripheral parts of the vertebrate nervous system including the autonomic innervation of the digestive tract (Mullen et al., 1992). The epitope of this antibody (see **Table 1**) matched at least seven nucleotide sequences (average length >1,000 bp) belonging to the O. vulgaris transcriptome available to us. In particular, we identified it by BLAST as the RNA binding protein fox-1 homolog 3 (RBFOX3). To the best of our knowledge, RBFOX3 gene encodes a member of the RNAbinding FOX protein family, which is involved in the regulation of alternative splicing of pre-mRNA. It has an RNA recognition motif domain, and is known to produce the neuronal nuclei (i.e., NeuN) antigen that has been widely used as a marker for post-mitotic neurons (see https://www.ncbi.nlm.nih.gov/gene/ 146713; see also Nikolic et al., 2013 ´ where is reported to identify neural cells in Helix).

In our experiments, NeuN antibody identified a population of cells in the octopus gastric ganglion. According to the original description of Bogoraze and Cazal (1946), the nerve cells forming the cortical layers (4 to 5, sensu Bogoraze and Cazal, 1946) are all of "ganglionic type (plasmo- or somatochromes), large in size, abundant in cytoplasm, rich in Nissl bodies, with vesicular and nucleolus nuclei" (Bogoraze and Cazal, 1946, p. 123; **Figure 4M**, upper row). Following the authors' description, there are no karyo-chrome cells expected in the octopus gastric ganglion (Bogoraze and Cazal, 1946). In addition, neural cells of the cortical layers are described as placed in a neuronal "lodge" formed by fibers and agglutinated neuroglia proposed to be part of the intricate glio-vascular network characterizing typical ganglia in octopus (see Bogoraze and Cazal, 1944, 1946, p. 123). Based on the NeuN, noradrenaline and GABA immunoreactivities we observed, it is suggested that our data match with the description provided by Bogoraze and Cazal. Interestingly, the intricate network seen for noradrenaline and GABA (**Figures 5I,J**) resembles the original drawings of the gastric ganglion histology (see Figure XIII in Bogoraze and Cazal, 1944).

It was not possible to investigate the glia-like cells in the gastric ganglion of octopus, as specific markers are not available for cephalopods. Imperadore et al. (2017) were unsuccessful using antibodies against vimentin and glial fibrillary acidic protein in octopus neural tissues; this contrasts with previous results (Cardone and Roots, 1990). Furthermore, transcripts for glial fibrillary acidic protein in O. bimaculoides genome or in O. vulgaris transcriptome do not seem to occur (Imperadore et al., 2017).