CRUSTACEAN BIOENERGETICS: MITOCHONDRIAL ADAPTIVE MOLECULAR RESPONSES TO FACE ENVIRONMENTAL CHALLENGES.
Crustaceans are one of the most diverse groups of invertebrate animals; the subphylum includes up to 17,635 species (De Grave et al. 2009). Some crustaceans have an ancient origin and a large evolutionary history based on limited paleontological evidence which is not well understood, not only because scientific efforts are limited, but also because of the highly diverse environments inhabited by these species.
To date, fossil evidence including the identification of a 520 million-y-old fossil of Ercaia, which was one of the earliest crustaceans ever found, suggests that crustaceans originated during the middle Cambrian (Chen et al. 2001). Marine crustaceans appeared during the Paleozoic, when some species such as the currently extinct Carpopenaeus spp., occurred (Rehm et al. 2011), then freshwater species, including the crayfish families such as Astacidae, Cambaridae. and Parastacidae, emerged much later, after the first terrestrial arthropods conquered land during the Silurian [443 to 419 million years ago (mya)], and are assumed to have their monophyletic origins during the Triassic period (185 to 22 mya) before Pangea split (Crandall et al. 2000). Finally, the existence of the first terrestrial crustaceans. Isopods, was confirmed by fossil record during the Cretaceous period (145 to 66 mya, Broly et al. 2013).
Although some million-year gaps make unclear the freshwater-species origin, the ability of crustaceans to successfully invade new environments is clear (Glenner et al. 2006). The abilities reproduction in both aerial and aquatic environments point toward the presence of adapted subcellular components, such as mitochondria. These organelles appeared on earth much earlier than animals, around 2.000 mya (during the Precambrian). when changing levels of atmospheric oxygen seem to have driven the mitochondrial evolution in species whose energetic needs promoted the origin of new molecules and highly specialized metabolic pathways (Lane 2002).
Crustacean diversity is represented through their highly specialized morphological and physiological characteristics. Some species may be found in highly extreme environments, where extensive physiological adaptations are required to ensure survival. For example, the amphipods Apherusa glacialis and Gammarus wilkitzkii, which inhabit the Arctic sea ice, possess physiological strategies to survive low temperatures, cope with food scarcity, and their metabolic rates are adapted to an energy-saving lifestyle (Poltermann et al. 2000. Werner & Auel 2005).
Parasitic crustaceans are also interesting species as they can infect a wide range of marine and freshwater invertebrates and vertebrates, mainly fish (Smit et al. 2014). Copepods, isopods. and brachyuran species show an amazing variety of adaptations, including the ability to redirect all the reproductive energy from a host through a process termed "host castration." which allows parasitic species to use and control the host energy, behavior, and physiology (Laffcrly & Kuris 2009). Of relevance also are the energetic mechanisms of low-oxygen tolerant species, such as the brine shrimp Artemia franciscana, whose quiescent embryos respond to long-term anoxia through down-regulation of metabolic processes and cessation of the transcription of mitochondrial proteins (Kwast & Hand 1996. Eads& Hand 2003). In addition, a number of various shrimp and lobster species have shown migratory behavior throughout their life cycle, which exposes them to highly variable environmental conditions (Dall et al. 1990).
At this point, bioenergetics, defined as the series of biological processes involved in energy flow and its chemical transformation, become central when studying mitochondrial genes and proteins, their function, and regulatory mechanisms to control energy expenditure, as well as the physiological response to environmental changes. Furthermore, it should be noted that mitochondria play also a central role in the generation of byproducts known as reactive oxygen species (ROS) and in their control through the activation of antioxidant systems and during the initiation of cell apoptosis. These processes are still scarcely understood in crustacean's mitochondria, and this knowledge will be helpful to researchers when trying to elucidate the abilities of crustaceans to regulate their energetic balance, which determines the degree of fitness and species perpetuation.
Fortunately, during the past decade, an increasing number of studies have focused on understanding the biology of crustaceans such as shrimp, lobsters and crabs, which are the most studied species because of their commercial values. In this review, we summarize the knowledge concerning crustacean bioenergetics, mitochondrial genes, and proteins and, specifically, the physiological response of mitochondria to environmental changes such as hypoxia. We also summarize current information regarding the transition permeability phenomenon and the presence of alternative enzymes in their respiratory chain or alternative sources and mechanisms to produce adenosine triphosphate (ATP).
CRUSTACEAN MITOGENOMES AND THE SHRIMP TRANSCRIPTOME
During the past three decades, the study of crustacean biology has increased considerably, and the use of novel techniques and tools has provided a better understanding of the biology of these species. Although only one crustacean nuclear genome data set has been published to date (the waterflea Daphnia pulex, Colbourne et al. 2011), the mitochondrial genomes (mitogenomes or mtDNAs) of several crustacean species reported to date have provided valuable information for establishing their evolutionary history, their phylogenetic relationships, and patterns of gene flow between species (Wilson et al. 2000).
To our knowledge, Artemia franciscana was the first crustacean species whose mtDNA was reported (Valverde et al. 1994). The total length of the A. franciscana mtDNA is 15.N22 base pairs (bp), a similar size to the mtDNA of insects such as the fly Drosophila yakuba (16,019 bp, Clary & Wolstenholme 1985) and the bee Apis metlifera (16,343 bp, Crozier & Crozier 1993). Some other crustacean mtDNAs have also been reported, including those of the waterflea Daphnia pulex (Crease 1999), the tiger shrimp Penaeus monodon (Wilson et al. 2000), the copepod Tigriopus japonicus (Machida et al. 2002), the Japanese blue crab Portunus trituberculatus (Yamauchi et al. 2003), and the freshwater crab Geothelphusa dehaani (Segawa & Aotsuka 2005). To date, there are more than a hundred reported mtDNAs from crustaceans, including 86 from Malacostracans, which is the largest of the six classes of crustaceans and includes extinct and living species, such as shrimp, lobsters, crabs, krill. mantis, and crayfish (Shen et al. 2015). The mtDNA sequences of all these species include the 13 mitochondrial-eneoded proteins, although some genes appear in different arrangements or include unusual characteristics, which are species specific (Kilpert & Podsiadlowski 2006). These 13 mitochondrial proteins encode subunits constituting the mitochondrial multimeric complexes involved in the electron transport chain and oxidative phosphorylation (Staton et al. 1997, Shen et al. 2007). Among crustaceans, shrimp, because of its commercial importance, are some of the most widely studied species. Particularly, the Pacific whiteleg shrimp Litopenaeus vannamei, whose mtDNA was reported in 2007 by Shen et al, will be considered as a key species in this review.
Besides mitogenomes. during the last 10y, molecular biology advances have led to the sequencing of transcriptomes (nuclear and mitochondrial transcripts) of an increasing number of crustaceans, which also provide a major source of information to better understand the mitochondrial machinery.
It is well known that in addition to the mitochondrial-eneoded proteins, most mitochondrial multimeric complexes from the respiratory chain require a series of nuclear-encoded proteins, which are imported from the cytosol to the mitochondria, to assemble the active complex.
In this review, the expressed sequence tags of some nuclear transcripts encoding mitochondrial proteins were selected from the Litopenaeus vannamei transcriptome database (Ghaffari et al. 2014) and their deduced proteins analyzed. Table 1 shows a summary of some of those shrimp proteins that may play central roles within the mitochondria, including those from metabolic pathways such as the Krebs cycle, the electron transport chain, and ATP synthesis.
All the transcripts analyzed in Table 1 were blasted against the National Center for Biotechnology Information (NCBI) database using BLASTx and compared with protein sequences from other animal species available in the GenBank database. No mtDNA-encoded proteins were included in the analysis because they were previously reported by Shen et al. (2007). The analyzed proteins include the complex I or NADH dehydrogenase subunits, the A and B subunits of complex II or succinate dehydrogenase, and various subunits of complex III or cytochrome bcl. The analyzed subunits of complex IV or cytochrome c oxidase (COX) included only proteins whose transcripts have not been reported to date. Previously, some of the full-length COX transcripts were sequenced and characterized such as: COX4 (GenBank accession no. JQ828862.1), COX5A (GenBank accession no. JQ839284.1), COX5B (GenBank accession no. JQ952564.1) (Jimenez-Gutierrez et al. 2013), and COX6A (GenBank accession no. KF906252.1), COX6B (GenBank accession no. KF906253.1) and COX6C (GenBank accession no. KF906254.1) (Mendez-Romero 2014); also their genes expressions have been analyzed.
Many deduced proteins encoding complex V or ATP-synthasc subunits are included in Table 1 along with the nuclear encoded ATP9 (GenBank accession no. EU194608.1, Muhlia-Almazan et al. 2008), and the main nuclear-encoded subunits from the F, domain of ATP synthase such as subunits ATP[alpha] (GenBank accession no. GQ848643.3), ATP[beta] (GenBank accession no. GQ848644.1), ATPy (GenBank accession no. HM036579.1), ATP[delta] (GenBank accession no. HM036580.1), and ATP[epsilon] (GenBank accession no. HM036581.1, Martinez-Cruz et al. 2011. Martinez-Cruz et al. 2015). The previously reported ATPase inhibitory protein IF, (GenBank accession no. KF306266.1) was not included in Table 1, although it is a central regulatory protein which has been associated with the ATP hydrolysis regulatory mechanism of shrimp mitochondria (Faccenda & Campanella 2012, Chimeo et al. 2015).
Table 1 also includes other important deduced proteins found in the shrimp transcriptome, such as those previously reported in mammals to act as regulatory proteins and those that may play a role during the permeability transition (PT) of the mitochondrial membrane; these include voltage dependent anion channel, cyclophilin D, the phosphate carrier protein, and the adenine nucleotide translocator (Bernardi 2013).
Data in Table 1 contain a comparison of the shrimp-deduced proteins with the homologous proteins from other marine invertebrates, including crustaceans as the copepods Tigriopus californicus and Lepeophtheirus salmonis, the crayfish Marsupenaeus japonicus, the water flea Daphnia pulex, the crab Eriocheir sinensis, and several species of molluscs (Lingula anatina, Crassostrea gigas, Biomphalaria glabrata, and Aplysia californica). In addition, it is important to point out the high similarities detected between crustacean and insect protein sequences. This finding is in agreement with the analysis previously described in phylogenetic studies from different mitochondrial shrimp proteins which suggests the existence of a shared common ancestor, Pancrustacea, of Crustacea and insects (Regier et al. 2010, Chimeo et al. 2015, Martinez-Cruz et al. 2015).
MITOCHONDRIAL RESPONSES TO OXIDATIVE STRESS AND ROS PRODUCTION IN CRUSTACEANS
Mitochondria, considered the powerhouse of the cell, produce ATP from adenosine diphosphate and inorganic phosphate through a coupled mechanism. The ATP synthesis involves a series of electron transport reactions in the inner mitochondrial membrane, where oxygen is an essential component to facilitate the oxidation of substrates that ultimately allows ATP production (Mitchell 1966, Boyer 1997).
In the aquatic environment, oxygen is less available to organisms than it is in the air. During shrimp adult and early stages, at the open sea, small variations are detected in the water oxygen levels; however, after postlarvae migration to low deep refuge zones as estuaries, or even in the culture ponds, oxygen levels may vary continuously from normoxia during daytime to hypoxia at night (Puente-Carreon 2009, Fig. 1).
Thus, water-breathers have fixed adaptive mechanisms to obtain this valuable molecule, even in those extreme environments with very low dissolved oxygen concentrations, and maintain the mitochondrial machinery functions. Indeed, the availability of molecular oxygen determines the adaptive ability of some species to face the stress produced by low [O.sub.2] concentrations, and the damage that increased ROS produces (Guzy & Schumacker 2006, Welker et al. 2013).
The Effect of Hypoxia on the Mitochondrial Function of Crustaceans
In marine ecosystems, the transitory fluctuations in oxygen concentration may lead to death in unadapted marine species (Vaquer-Sunyer & Duarte 2008). The physiological responses of individual species to changes in the environmental oxygen concentration may help to distinguish those marine animals that tolerate anoxia/hypoxia from those which are sensitive.
Whereas sensitive organisms use energy compensatory pathways, tolerant species instead use energy conservatory mechanisms to maintain their energy balance (Gorr et al. 2006). In the presence of low oxygen levels, unadapted or hypoxia-sensitive organisms display physiological responses that stimulate specific metabolic pathways to increase ATP production, and under such condition, there is a marked shift from aerobic to anaerobic metabolism. Hypoxia-sensitive animals also show a quick decay in cellular ATP concentration, causing activation of proteases, an uncontrolled rise in the concentration of calcium ions, continuous membrane depolarization, and cell death under low oxygen conditions (Lutz & Milton 2004, Nilsson & Lutz 2004). On the other hand, animals adapted to hypoxia maintain normal ATP levels by entering a reversible hypometabolic state, in which pathways consuming ATP are repressed in the presence of restricted oxygen availability (Boutilier 2001, Hochachka & Lutz 2001, Storey 2004). In addition, some of these species may store large glycogen reserves in their tissues, are able to activate powerful antioxidant systems, and may possess mitochondrial enzymes, such as alternative oxidases (AOX) and uncoupling proteins (UCP) that may help to control ROS formation (Lutz & Nilsson 1997, Hermes-Lima & Zenteno-Savin 2002, Sokolova & Sokolov 2005, Abele et al. 2007).
Various shrimp species are thought to be hypoxia-sensitive because low oxygen levels may affect their mitochondrial functions (Wu et al. 2002, Sun et al. 2015); however, some anoxia-adapted species, such as Artemia franciscana, have developed interesting strategies to survive prolonged periods of anoxia, many of which are, presumably, tightly related to their mitochondrial adaptations (Menze et al. 2005).
Recent studies have focused on the mitochondrial responses of crustaceans after exposure to hypoxia and re-oxygenation events. At the transcriptional level, hypoxia significantly affects the expression of various nuclear-encoded mitochondrial proteins (Jimenez-Gutierrez et al. 2014, Martinez-Cruz et al. 2015). The hypoxia-inducible factor (HIF) has been identified as the main regulator of the transcriptional responses to hypoxia in mammalian mitochondria. This transcription factor with a heterodimeric structure comprised an oxygen-regulated a subunit (HIF-[alpha]), and a constitutively expressed [beta] subunit (Semenza 2000) is translocated to the nucleus during hypoxia, where it binds to specific DNA sequences to regulate a large number of genes implicated in energetic metabolism, including 6-phosphofructo-2-kinase, fructose-2,6-biphosphatase 3 and 4, and COX subunits 4-1 (Minchenko et al. 2003, Fukuda et al. 2007). In crustaceans. HIF has been characterized only in a few species, including Litopenaeus vannamei, Palaemonetes pugio, Daphnia magna, and Callinectes sapidus, and its ability to control a variety of cellular and systematic homeostatic responses to hypoxic stress has been partially confirmed (Gorr et al. 2004, Li & Brouwer 2007, Sonanez-Organis et al. 2009, Hardy et al. 2012).
Recently, new high-throughput sequencing technologies have been used to examine the effects of hypoxia on the transcriptome of some crustaceans, including the red swamp crayfish (Procambarus clarkii), the oriental river prawn (Macrobrachium nipponense), and the white shrimp (Litopenaeus vannamei). The results indicated an increase in the expression of genes encoding proteins implicated in the mobilization of carbohydrates and lipids, downregulation of genes involved in the protein degradation machinery, and no major changes in genes encoding proteins of the translational machinery after exposure to hypoxia (Shen et al. 2014, Johnson et al. 2015, Sun et al. 2015).
The transcripts of some nuclear-encoded proteins from the mitochondrial respiratory chain of the shrimp Litopenaeus vannamei are known to be downregulated by hypoxia, including some proteins that compose the Frcatalytic domain of the ATP synthase (Martinez-Cruz et al. 2015), the regulatory [IF.sub.1] mitochondrial protein that inhibits ATPase activity (Chimeo et al. 2015), and some nuclear-encoded subunits of the COX complex (Jimenez-Gutierrez et al. 2013).
In addition to transcriptional changes, several other biochemical and physiological alterations have been observed in shrimp-isolated mitochondria in response to hypoxia/re-oxygenation. The synthesis of mitochondrial proteins, mainly those that are part of the COX and ATP synthase complexes, were significantly affected by the dissolved oxygen concentration of seawater (Martinez-Cruz et al. 2012, Jimenez-Gutierrez et al. 2014), and lower levels of proteins such as ATP[beta] and COX1 (the catalytic subunits of mitochondrial complexes IV and V) have been observed under hypoxic conditions when compared with normoxia. Also, COX activity and the oxygen consumption of muscle-isolated mitochondria significantly decrease under hypoxia (Jimenez-Gutierrez et al. 2014).
Opposite to other enzymes, the mitochondrial ATPase activity of Litopenaeus vannamei increased during hypoxia (Martinez-Cruz et al. 2012), an adaptation that was suggested to maintain the membrane potential and cellular homeostasis, as previously observed in vertebrate species. It has been proposed that in shrimp muscle, the presence of arginine-phosphate may act as a buffer to maintain the tissue ATP levels during hypoxia. As previously mentioned, shrimp may face daily hypoxia/re-oxygenation cycles throughout their lifetime, and these events may be analogous to the oxidative stress that ischemia/reperfusion episodes can provoke in well-studied vertebrate models (Abele et al. 2012). In fact, re-oxygenation, which is the abrupt increase in the oxygen concentration after hypoxia, seriously affects mitochondrial function in invertebrates because of an increase in ROS production, which may provoke profound cellular damage (Hermes-Lima & Zenteno-Savin 2002, Solaini & Harris 2005). Previous studies have confirmed that shrimp rapidly respond to increasing oxygen levels after hypoxia by reducing ATPase activity and ATP levels. Furthermore, the activities of mitochondrial enzymes, such as citrate-synthase and COX, which are downregulated during hypoxia, increase abruptly (Martinez-Cruz et al. 2012, Jimenez-Gutierrez et al. 2014). At re-oxygenation. white shrimp are able to restore their mitochondrial functions to those observed at normoxia; however, recent findings suggest that many of the biochemical responses observed at re-oxygenation as the L-lactate levels in plasma may return to the previous levels after 12-24 h of re-oxygenation, as observed in L. vannamei (Chimeo et al., submitted).
These findings suggest that shrimp mitochondria have the ability to conform to changes in environmental oxygen availability because their oxygen consumption rate increases/decreases in direct relation with the dissolved oxygen levels (data not shown), supporting the hypothesis that shrimp species are not as sensitive to hypoxia as previously suggested. Future studies, however, are required to fully understand, at the organelle level, how shrimp mitochondrial enzymes deal with continuous fluctuations in the dissolved oxygen concentration of seawater and to elucidate the strategies these species use to control mitochondrial ROS production under these conditions, besides the activation of their antioxidant system.
ROS Production and Mitochondrial Responses to Oxidative Stress in Crustaceans
Oxidative stress results in an increase in the production of ROS, mainly as byproducts during mitochondrial electron transport. The oxygen radical superoxide ([O.sup.*-.sub.2]) and hydrogen peroxide ([H.sub.2][O.sub.2]) are commonly produced during normal oxidative metabolism. It has been established that an excessive production of ROS has been associated with a range of pathological conditions that may result in tissue and cell damage (Kowaltowski et al. 2009). Those aquatic species that cyclically encounter broad changes in dissolved oxygen concentration may cope with the damage caused to cell proteins, lipids, and nucleic acids by an overproduction of ROS (Lawniczak et al. 2013).
Some crustaceans have shown an increase in lipids peroxidation in response to oxidative stress brought on by episodes of hypoxia/anoxia. In 2005, de Oliveira et al. used the lipids peroxidation assay (thiobarbituric acid reactive substances, TBARS) to demonstrate that lipid peroxidation significantly increases in the gills of the estuarine crab Chasmagnathus granulate during anoxia recovery. In addition, during air exposure (when intertidal crabs are not in seawater), lipids peroxidation occurs in various crustacean tissues, including the hepatopancreas, muscle, anterior and posterior gills of crab species such as Callinectes danae and Callinectes ornatus as a response to the oxidative damage caused by ROS (Freire et al. 2011).
In addition to lipid peroxidation, protein carbonyl groups produced as a result of oxidative protein damage have been detected in hepatopancreas, muscle, and gills of crustaceans such as the stone crab (Paralomis granulosa) after 24 h of air exposure (Romero et al. 2007); thus, damage signals in lipids and proteins are a consequence of ROS accumulation in the mitochondria after hypoxia/anoxia. Nevertheless, certain organs in some marine invertebrates, such as the hepatopancreas of the snail Littorina littorea (Hermes-Lima & Zenteno-Savin 2002), and muscle, hepatopancreas, and gills of the white shrimp Litopenaeus vannamei (Zenteno-Savin et al. 2006), are unaffected by anoxia and do not demonstrate lipid peroxidation after hypoxia. This lack of lipid and protein damage by ROS has been suggested to be organ-dependent and may be related to several factors, including the duration of hypoxia/anoxia/re-oxygenation exposure, the antioxidant capacity of the specific organ/tissue, and/or species-specific mitochondrial mechanisms that may control ROS generation.
In crustaceans, the increase in mitochondrial ROS production during hypoxia/re-oxygenation acts as a signal that triggers cellular responses that initially include activation of the antioxidant systems, both enzymatic (superoxide dismutase, catalase, glutathione-peroxidase, and peroxiredoxin) and non-enzymatic (vitamins C, E, thiamine, and glutathione) molecules that are known to be induced and modulated by many factors, including oxygen deprivation, the presence of heavy metal ions, pathogen infection, and ammonia stress. The extent of activation of these antioxidant systems varies throughout the life cycle in crustaceans (Aispuro-Hernandez et al. 2008, Parrilla-Taylor & Zenteno-Savin 2011, Abele et al. 2012). At this point, a new central question emerges; besides the activation of antioxidant systems, do crustacean mitochondria have another mechanism(s) to deal with the increased ROS production stimulated by hypoxic/re-oxygenation conditions?
The Mitochondrial Uncoupling Mechanisms of Crustaceans and Other Marine Invertebrates
The mitochondrial electrochemical gradient, or proton motive force ([DELTA][p.sub.m]), is generated through the electron transport chain and used by the ATP synthase to produce ATP. This [DELTA][p.sub.m] involves two components: an electrical constituent denoted as the mitochondrial transmembrane potential ([DELTA][PSI]) and a chemical constituent, the pH difference across the mitochondrial inner membrane (MIM) (Mitchell 1966, Sanderson et al. 2013).
Besides the [DELTA][PSI] role in ATP synthesis, it is also involved in the production of ROS because an increased [DELTA][PSI] is known to slow electron transfer among the enzymes in the respiratory chain; intermediates react allowing a single electron reduction of oxygen, this produces an electron scape and superoxide anion formation, which is then converted to other ROS (Murphy 2009).
Within the mitochondria, [DELTA][PSI] tends to increase during nutrient oxidation and decreases during oxidative phosphorylation or mitochondrial uncoupling. This allows the uncoupling of ATP synthesis from the proton electrochemical gradient via a proton leak into the mitochondrial matrix, promoting a decreased [DELTA][PSI] without ATP production (Slocinska et al. 2011). To date, various studies have confirmed that ROS production can be controlled by regulating [DELTA][PSI] (Drose & Brandt 2012), and the mitochondrial uncoupling mechanisms have been suggested to prevent a high [DELTA][PSI] (Kowaltowski et al. 2009).
The uncoupling mechanisms that may be linked to the high resistance of some species to reoxygenation-derived oxidative stress by reducing ROS production include: 1) proton sinks that involve the UCP, the bacterial mitochondrial unspecific channels, or the PT pore (PTP) of mammals; and 2) the non-pumping alternative redox enzymes including a NADH-dehydrogenase type 2 (NDH2), the mitochondrial glycerol-3-phosphate-dehydrogenase (mtGPDH), and the AOX, each as an additional participant in a branched respiratory chain (Kadenbach 2003, Lesser 2006, Guerrero-Castillo et al. 2011).
The UCP are mitochondrial carrier proteins located in the inner membrane, and considered uncouplers, because these proteins are capable of dissociating ATP synthesis from the respiratory chain by dissipating the proton gradient. This protein family includes a core group of five mammalian UCP variants (UCP 1-5), which are differentially expressed in a tissue-specific manner (Chan et al. 2006, Bermejo-Nogales et al. 2014).
In the last decade, the number of studies of invertebrate UCP has risen. Various studies have reported at least two UCP--UCP4 and UCP5--in the mitochondria of invertebrates, including parasites such as the nematode Caenorhabditis elegans (UCP4), insects such as the fruit fly Drosophila melanogaster (UCP4), the blood sucking insect Rhodnius prolixus (UCP4), and the cockroach Gromphadorhina cocquereliana. These studies have suggested that UCP4 is involved in regulating lipid metabolism and/or in protecting against oxidative stress (Slocinska et al. 2011. Ji et al. 2012, Alves-Bezerra et al. 2014. Da-Re et al. 2014). To date, the eastern oyster Crassostrea virginica (UCP5) is the only marine mollusc in which the presence of UCP has been demonstrated (Kern et al. 2009). Recently, our research group has confirmed the existence of UCP4 and UCP5 in the white shrimp mitochondria (Mendez-Romero 2016).
The Mitochondrial Permeability Transition Enigma in Crustaceans
The physiological uncoupling ability of mitochondria is closely related to the inner membrane PT. This phenomenon, which results from the opening of the mitochondrial unspecific channel in bacteria, has been confirmed in mammals and in some yeasts such as Saccharomyces cerevisiae and Debaryomyces hansenii (Cabrera-Orefice et al. 2010, Guerrero-Castillo et al. 2011). In mammalian mitochondria, PT reflects an increased permeability of the MIM that results from stress conditions which promote an intracellular calcium imbalance accompanied by elevated phosphate concentrations and adenine nucleotide reduction (Halestrap et al. 1998). The long-lasting PT enables the free passage of protons and molecules less than 1.5 kDa in size into the mitochondrial matrix; as a result, the proton barrier disappears, and the [DELTA][p.sub.m] is no longer preserved (Halestrap 2009. Bernardi 2013).
It has been well established that mitochondrial PT can be triggered by anoxia/hypoxia and ischemia/reperfusion events, as well as pathological conditions that may induce apoptosis (Bernardi et al. 2006, Menze et al. 2010). This response was first assumed to be mediated by the opening of a multiprotein channel or pore, which induces mitochondrial swelling, membrane depolarization, uncoupled respiration, and if prolonged, leads to the rupture of the outer mitochondrial membrane, the release of cytochrome c, and, ultimately, apoptosis of the cell (Kroemer et al. 2007, Giorgio et al. 2013).
To date, scarce information is available about the ability of crustacean mitochondria to undergo PT (Menze et al. 2010). Early studies in the mitochondria from Artemia franciscana embryos suggested the existence of the three main proteins forming the mitochondrial pore: the voltage dependent-anion channel (VDAC), the adenine nucleotide translocator (ANT), and the mitochondrial cyclophilin D (Cyp D) as it was first suggested for the mammalian mitochondrial pore (Halestrap & Davidson 1990, Menze et al. 2005); however, the high calcium loads that promote those responses observed in mammalian mitochondria were not detected in Artemia; therefore, the existence of a non-calcium regulated pore in this species was suggested (Menze et al. 2005). Mitochondria from other crustaceans including the ghost shrimp Lepidophthalmus louisianensis (Holman & Hand 2009) and the northern shrimp Crangon crangon and Palaemon serratus (Konrad et al. 2012) have been studied, and the compiled evidence suggests that crustacean mitochondria lack the ability to induce a PT in response to a cellular calcium increase.
In 2010, Menze et al., suggested this non-calcium-regulated pore may be specific to arthropods because no PT was detected in the mitochondrial membrane of the oyster Crassostrea virginica (Sokolova et al. 2004); however, von Stockum et al. (2015) recently reported that, similar to mammals, the mitochondria from Drosophila melanogaster have a [Ca.sup.2+]-induced [Ca.sup.2+] release channel (mCrC) that shares regulatory features with the PTP and opens in a [Ca.sup.2+]-induced manner. To our knowledge, these findings suggest that in crustaceans, mitochondria PT may not be regulated, or at least not regulated by [Ca.sup.2+], but instead by another ion (Krumschnabel et al. 2005).
In fact, inhibition or resistance of crustacean mitochondria to induced PT may represent a mechanism for depressing apoptosis, as suggested by Menze et al. (2010); however, the regulation of apoptosis in crustaceans is currently not well understood, and future experiments are required to more fully examine the function of the mitochondrial PTP in crustaceans and to ultimately support or reject this hypothesis.
The Alternative Mitochondrial Enzymes of Marine Invertebrates
An additional mitochondrial uncoupling strategy may involve the participation of various alternative respiratory enzymes. which may be present in the mitochondrial apparatus. These non-pumping enzymes are part of a branched respiratory chain and transfer electrons from one enzyme to the next one without pumping protons from the mitochondrial matrix to the inter-membrane space, thereby differing from the classical transport chain, and promoting a decreased [DELTA][PSI] and ROS generation (McDonald et al. 2009).
To date, an alternative type-2 NADH dehydrogenase (NADH2), a mitochondrial glycerol-3-phosphate dehydrogenase (mtGPDH), and an AOX are the three enzymes reported as part of the mitochondrial branched respiratory chains of mammals, yeast, and marine invertebrates (Abele et al. 2007, Mracek et al. 2013). The NADH2 serves as an alternative to the multisubunit respiratory complex I, and catalyzes the transfer of electrons from NADH to ubiquinone in the mitochondrial respiratory chain. In yeast, the NADH2 oxidizes NADH on the matrix side and reduces ubiquinone to maintain mitochondrial NADH/NAD(+) homeostasis without generating an electrochemical gradient (Feng et al. 2012). This enzyme may be found mainly in bacteria, yeast, fungi, and Arabidopsis thaliana, but not in mammals or arthropods (Iwata et al. 2012). There are currently no reports of the presence of these enzymes in crustaceans or other invertebrate marine organisms.
The mtGPDH is the smallest protein found in the mammalian respiratory chain (74 kDa). This enzyme is located on the outer side of the MIM, is FAD-dependent, and has been suggested to be a major source of ROS production in the mitochondrial intermembrane space. The function of mtGPDH has largely been discussed in mammals and invertebrates such as the fruit fly Drosophda melanogaster; the enzyme may act as part of the mitochondrial electron transport chain by transferring electrons to ubiquinone and acting as an uncoupling mechanism susceptible to proton leak (Miwa et al. 2003, Vrbacky et al. 2007, Carmon & Maclntyre 2010). Among invertebrates, an mtGPDH was localized in the mitochondria of the parasites Trypanosoma brucei and Leishmania major (Skodova et al. 2013), but to date there are no reports confirming the existence of this enzyme in crustaceans.
The AOX, also known as ubiquinol oxidase, catalyzes the oxidation of ubiquinol and the reduction of four electrons from molecular oxygen to water; it is alternative to the COX function but without pumping protons to the intermembrane space, insensitive to cyanide, and is located in the MIM. In mammals, AOX prevents an over-reduction of ubiquinone, and a consequent overproduction of superoxide (El-Khoury et al. 2013). In marine invertebrates, such as annelid worms, sipunculid worms, and molluscs, the presence of an AOX has been described; nevertheless, to date there are no reports of this enzyme in crustaceans (McDonald et al. 2009).
In 2007, Abele et al., suggested that when the mitochondrial respiratory rate of some marine invertebrate hypoxia-tolerant species, including the bivalve Arctica islandica, decreases under disadvantageous conditions, the mitochondrial AOX may help to maintain the mitochondrial respiratory rate without disturbing the energetic balance of cells. In addition, AOX may also exhibit an antioxidant function by minimizing ROS formation. Recently, Sussarellu et al. (2012) detected a significant increase in expression of the AOX transcripts of the Pacific giant oyster Crassostrea gigas during re-oxygenation, and confirmed this increase as a protective mechanism against the abrupt increase in ROS production.
Crustacean mitochondria possess some peculiarities when compared with other related groups. There are presently no published studies examining mitochondrial uncoupling mechanisms in crustaceans. So far, new evidence resulting from the identification of specific mitochondrial transcripts and proteins, and the determination of mitochondrial responses from various crustacean species, suggests the existence of putative AOX (data not shown) and UCP in decapods (Mendez-Romero 2016). Further research in this area is needed to confirm the functionality of these enzymes and proteins, its participation in the mitochondrial antioxidant defense, and to correlate its function with the ability of some crustacean species to tolerate hypoxia/anoxia and avoid ROS production.
CRUSTACEAN PHYSIOLOGICAL PROCESSES, IONS TRANSPORT, AND ATP PRODUCTION
Although a considerable number of studies have described in great detail, the transport of specific ions such as calcium and sodium in the mitochondria of vertebrate cells, studies on these ions transport mechanisms in crustaceans are still limited. In vertebrates, calcium plays essential roles in regulating the mitochondrial volume and the regulatory activities of some calcium-dependent enzymes in the Krebs cycle. In fact, as previously discussed in The Mitochondrial Permeability Transition Enigma in Crustaceans, various calcium-dependent transport mechanisms may directly influence mitochondrial functions and cell death (Bernardi 1999, Campanella et al. 2004).
In crustaceans, calcium is not only required to complete basic cellular functions but has also been shown to be essential for meeting energetic demands during growth and molting (Zilli et al. 2003, 2007). The role of calcium in crustacean hepato-pancreatic cells has also been studied, and calcium has been determined to participate in regulating the concentration of heavy metals, including copper, zinc, iron, and magnesium, which play important roles in enzymes activation and in the synthesis of respiratory pigments such as hemocyanin. At higher concentrations, such metal ions become toxic, and a regulatory mechanism involving mitochondrial heavy-metal sequestration has been suggested in the mitochondria of epithelial hepatopancreatic cells from species such as the American lobster Homarus americanus. It has been proposed that this mechanism involves a calcium-heavy metal uniporter uptake that transports divalent cations from the cytoplasm to the mitochondria, where an insoluble nontoxic precipitate is accumulated (Chavez-Crooker et al. 2001, 2002, Ahearn et al. 2004).
To date, there is no information about the mitochondrial carrier proteins involved in the [Ca.sup.2+] flux of crustaceans; however, new published information about the transcriptome of species such as Litopenaeus vannamei (Ghaffari et al. 2014) confirmed the existence of various expressed sequence tags encoding mitochondrial proteins that participate in the calcium in- and out-fluxes. The existence of the mitochondrial [Ca.sup.2+] uniporter, which dynamically buffers physiological [Ca.sup.2+] concentrations in mitochondria, has been confirmed in this shrimp species (data not shown); however, to date there is no information that confirms the existence of proteins such as calcium antiporters either 2[N.sub.a+] or 2[H.sub.+] in crustaceans. Future studies are needed to better understand the extensive ability of shrimp mitochondria to uptake high calcium concentrations without suffering the mitochondrial PT phenomenon as it happens in all other species mitochondria.
ATP Production from Sulfide
In the 1930s it was first reported that some organisms are able to survive in the presence of sulfide, despite the fact that environments with high sulfide contents also have low dissolved oxygen concentrations in water, thus promoting anoxic or hypoxic marine conditions. To survive under these harsh environmental conditions, organisms have developed adaptive strategies, including the ability to cover their bodies and internal epithelial mucosa with sulfide-oxidizing bacteria (Grieshaber & Volkel 1998). In addition, early studies have suggested that mitochondria present in gills of the gutless clam Solemya reidi may convert sulfide into less toxic compounds and produce energy in a coupled reaction to synthesize ATP (Powell & Somero 1986).
Later, O'Brien and Vetter (1990) suggested that the mitochondrial electron transport chain complexes contribute to ATP production from sulfide and estimated that mitochondria of Solemya reidi produce approximately 2.0-3.2 ATP molecules per sulfide molecule. More than two decades later, Hildebrandt and Grieshaber (2008) proposed a mechanism to explain mitochondrial oxidation of sulfide in the lugworm Arenicola marina, which produces thiosulfate, a less toxic compound, and the simultaneous electron transfer from sulfide oxidation to the respiratory chain complexes III and IV. During this process, for each pair of electrons, six protons are translocated from the intermembrane space to the mitochondrial matrix and 1.5 ATP molecules are synthesized (Hinkle 2005).
Presently, three enzymes are known to be involved in the sulfide conversion and ATP production. The first enzyme is sulfide:quinone oxidoreductase, which is located in the MIM where it oxidizes sulfide ([H.sub.2]S) to persulfide (R-SSH) and transfers two electrons to the ubiquinone pool, as described by Theissen and Martin (2008). The second and third enzymes--a sulfur dioxygenase that achieves four-electron oxidation of one persulfide molecule to sulfite in the presence of oxygen and water, and a sulfur transferase, which transfers a sulfite to persulfide, resulting in the production of thiosulfate--are located in the mitochondrial matrix (Hildebrandt & Grieshaber 2008, Fig. 2).
Some additional studies have focused on the ability of species to tolerate high-sulfide concentrations, and the use of sulfide as a substrate to produce ATP have been found in crustaceans such as the shrimp species Calocaris macandreae, Callianassa subterranea, and Jaxea nocturna (Johns et al. 1997). In 2001, Bourgeois and Felder confirmed that mitochondria isolated from the ghost shrimp Lepidophthalmus louisianensis under high-sulfide and hypoxic conditions are able to synthesize ATP at a rate of 0.474-14.750 nmol ATP/min/mg protein. Furthermore, the thiosulfate levels increased in the hemolymph of these organisms during exposure to anoxic-sulfide water. Therefore, the authors suggested that ATP production from thiosulfate occurs in mitochondria through the electromotive force, which is generated by the electron/proton donor reaction between sulfide and thiosulfate (Bourgeois & Felder 2001). It seems that, in certain species, prolonged exposure to sulfide has allowed mitochondria to adapt their functions to include the production of ATP from this alternative energy source.
The central role of mitochondria in cellular bioenergetics has been reviewed extensively elsewhere. This organelle directly uses oxygen to produce energy, and it inevitably produces toxic byproducts known as ROS. Although previous information provides some evidence on the way crustaceans have adapted to hypoxia, early studies focused on understanding the mitochondrial functions of these species seem to indicate diverse specific regulatory responses, which may lead future research to better understand their abilities to surpass the marine environment continuous variations, maintaining their bio-energetic equilibrium.
As previously discussed, the Pacific whiteleg shrimp, now suggested as a hypoxia-tolerant species, may enter into a reversible hypometabolic state when oxygen availability is low. At this condition, the genie expression of specific nuclear-encoded mitochondrial subunits from the respiratory chain is down-regulated and the hydrolytic activity of central enzymes as the mitochondrial ATPase is regulated through an [IF.sub.1] protein that may control ATP expenditure and the ATP levels of shrimp mitochondria decrease as a consequence of a prolonged lactate production even in the presence of oxygen during re-oxygenation. Under oxidative stress conditions, the antioxidant system of shrimp is well known to rapidly respond to the increasing ROS production; however, the existence and functionality of mitochondrial uncoupling mechanisms such as alternative enzymes and UCPs, are being studied and their ability to protect crustacean cells remains to be confirmed. In addition, it remains to be determined whether the existence and function of uncoupling mechanisms in mitochondria confer the ability to shrimp to efficiently face hypoxia/re-oxygenation cycles. In some other well-studied animal models, the ability of mitochondria to uncouple has been associated to the inner membrane PT. Nevertheless, previous studies suggest that crustacean mitochondria lack the capacity to undergo PT; this may also explain the resistance of shrimp mitochondria to hypoxia/re-oxygenation damage.
Several central questions regarding the mitochondrial functions in crustaceans remain unanswered. Does the PT phenomenon exist in crustacean mitochondria? What induces or regulates the PT in the inner mitochondrial membrane of crustaceans? How do shrimp mitochondria regulate calcium transport? Does hypoxia affect the calcium in- and out-flow in shrimp mitochondria? As many mitochondrial enzymes are multimeric complexes that participate as a whole in the energy-producing pathways, a central task to solve is the identification of all those proteins and subunits forming part of complexes, and regulatory mechanisms in crustaceans because there is clear evidence that confirms the existence of species-specific proteins that may accomplish new regulatory functions.
AMA and OMC acknowledge support from Consejo Nacional de Ciencia y Tecnologia (CON ACyT) and graduate scholarships to CC and CMR. This work was supported by Consejo Nacional de Ciencia y Tecnologia (grant 241670).
Abele, D., E. Philipp, P. M. Gonzalez & S. Puntarulo. 2007. Marine invertebrate mitochondria and oxidative stress. Front. Biosci. 12:933-946.
Abele, D., J. P. Vazquez-Medina & T. Zenteno-Savin. 2012. Oxidative stress in aquatic ecosystems. UK: Wiley-Blackwell. 548 pp.
Ahearn, G. A., P. K. Mandal & A. Mandal. 2004. Mechanisms of heavy-metal sequestration and detoxification in crustaceans: a review. J. Comp. Physiol. B 174:439-152.
Aispuro-Hernandez, E., K. D. Garcia-Orozco, A. Muhlia-Almazan. L. del Toro-Sanchez, R. M. Robles-Sanchez, J. Hernandez, G. Gonzalez-Aguilar, G. Yepiz-Plascencia & R. R. Sotelo-Mundo. 2008. Shrimp thioredoxin is a potent antioxidant protein. Comp. Biochem. Physiol. C Toxicol. Pharmacol. 148:94-99.
Alves-Bezerra, M., D. Cosentino-Gomes. L. P. Vieira, N. Rocco-Macchado, K. C. Gondim & J. R. Meyer- Fernandez. 2014. Identification of uncoupling protein 4 from the blood-sucking insect Rhodnius prolixus and its possible role on protection against oxidative stress. Insect Biochem. Mol. Biol. 50:24-33.
Bermejo-Nogales, A., J. A. Calduch-Giner & J. Perez-Sanchez. 2014. Tissue-specific gene expression and functional regulation of uncoupling protein 2 (UCP2) by hypoxia and nutrient availability in gilthead sea bream (Sparus aurata): implications on the physiological significance of UCP1-3 variants. Fish Physiol. Biochem. 40:715-762.
Bernardi, P. 1999. Mitochondrial transport of cations: channels, exchangers, and permeability transition. Physiol. Rev. 79:1127-1155.
Bernardi, P. 2013. The mitochondrial permeability transition pore: a mystery solved? Front. Physiol. 4:1-12.
Bernardi, P., A. Krauskopf, E. Basso, V. Petronilli, E. Blalchy-Dyson, F. Di Lisa & M. A. Forte. 2006. The mitochondrial permeability transition from in vitro artifact to disease target. FEBS J. 273:2077-2099.
Bourgeois. R. P. & D. L. Felder. 2001. Postexposure metabolic effects of sulfide and evidence of sulfide-based ATP production in callianassid ghost shrimp (Crustacea: Decapoda: Thalassimdea). J. Exp. Mar. Biol. Ecol. 263:105-121.
Boutilier, R. G. 2001. Mechanisms of cell survival in hypoxia and hypothermia. J. Exp. Biol. 204:3171-3181.
Boyer, P. D. 1997. The ATP synthase--a splendid molecular machine. Annu. Rev. Biochem. 66:1X1-149.
Broly, P., P. Deville & S. Maillet. 2013. The origin of terrestrial isopods (Crustacea: Isopoda: Oniscidea). Evol. Ecol. 27:461-416.
Cabrera-Orefice, A., S. Guerrero-Castillo, L. A. Luevano-Martinez, A. Pena & S. Uribe-Carvajal. 2010. Mitochondria from the salt-tolerant yeast Deharyomyces hansenii (halophilic organelles?). J. Bioenerg. Biomembr. 42:11-19.
Campanella, M., P. Pinton & R. Rizzuto. 2004. Mitochondrial [Ca.sup.2+] homeostasis in health and disease. Biol. Res. 37:653-660.
Carmon, A. & R. Maclntyre. 2010. The cc-glycerophosphate cycle in Drosophila melanogaster VI. Structure and evolution of enzyme paralogs in the genus Drosophila. J. Hered. 101:225-243.
Chan, S. L., D. Liu, G. A. Kyriazis, P. Bagsiyao, X. Ouyang & M. P. Mattson. 2006. Mitochondrial uncoupling protein-4 regulates calcium homeostasis and sensitivity to store depletion-induced apoptosis in neural cells. J. Biol. Chem. 281:37391-37403.
Chavez-Crooker, P., N. Garrido & G. A. Ahearn. 2001. Copper transport by lobster hepatopancreatic epithelial cells separated by centrifugal elutriation: measurements with the fluorescent dye Phen Green. J. Exp. Biol. 204:1433-1444.
Chavez-Crooker, P., N. Garrido & G. A. Ahearn. 2002. Copper transport by lobster (Hotnarus americanus) hepatopancreatic mitochondria. J. Exp. Biol. 205:405-413.
Chen, J. Y., J. Vannier & D. Y. Huang. 2001. The origin of crustaceans: new evidence from the early Cambrian of China. Proc. R. Soc. Lond. 268:2181-2187.
Chimeo, C, A. V. Fernandez-Gimenez, M. Campanella, O. Mendez-Romero & A. Muhlia-Almazan. 2015. The shrimp mitochondrial [F.sub.0][F.sub.1]-ATPase inhibitory factor 1 ([IF.sub.1]). J. Bioenerg. Biometnbr. 47:383-393.
Clary, D. O. & D. R. Wolstenholme. 1985. The mitochondrial DNA molecule of Drosophila yakuba: nucleotide sequence, gene organization, and genetic code. J. Mol. Evol. 22:252-271.
Colbourne, J. K., M. E. Pfrender, D. Gilbert, W. K. Thomas, A. Tucker, T. H. Oakley, S. Tokishita, A. Aerts, G. J. Arnold, M. K. Basu, D. J. Bauer, C. E. Caceres, L. Carmel, C. Casola, J. H. Choi, J. C. Detter. Q. Dong, S. Dusheyko. B. D. Eads, T. Frohlich. K. A. Geiler-Samerotte, D. Gerlach, P. Hatcher, S. Jogdeo, J. Krijgsveld, E. V. Kriventseva, D. Kultz, C. Laforsch. E. Lindquist, J. Lopez, J. R. Manak, J. Muller, J. Pangilinan, R. P. Patwardhan, S. Pitluck, E. J. Pritham, A. Rechtsteiner, M. Rho, I. B. Rogozin, O. Sakarya, A. Salamov. S. Schaack, H. Shapiro, Y. Shiga, C. Skalitzky, Z. Smith, A. Souvorov, W. Sung. Z. Tang, D. Tsuchiya, H. Tu, H. Vos, M. Wang, Y. I. Wolf, H. Yamagata. T. Yamada, Y. Ye, J. R. Shaw, J. Andrews. T. J. Crease. H. Tang, S. M. Lucas. H. M. Robertson, P. Bork. E. V. Koonin, E. M. Zdobnov, 1. V. Grigoriev, M. Lynch & J. L. Boore. 2011. The ecoresponsive genome of Daphniapulex. Science 331:555-561.
Crandall, K. A., D. J. Harris & J. W. Fetzner. 2000. The monophyletic origin of freshwater crayfish estimated from nuclear and mitochondrial DNA sequences. Proc. Biol. Sci. 267:1679-1686.
Crease, T. J. 1999. The complete sequence of the mitochondrial genome of Daphnia pulex (Cladocera: Crustacea). Gene 233:89-99.
Crozier, R. H. & Y. C. Crozier. 1993. The mitochondrial genome of the honeybee Apis mellifera: complete sequence and genome organization. Genetics 133:97-117.
Da-Re, C, C. De Pitta, M. A. Zordan, G. Teza, F. Nestola, M. Zeviani, R. Costa & P. Bernardi. 2014. UCP4C mediates uncoupled respiration in larvae of Drosophila melanogaster. EMBO Rep. 15:586-591.
Dall, W., B. J. Hill, P. C. Rothlisberg & D. J. Sharpies. 1990. The biology of Penaeidae, vol. 27. New York, NY: Academic Press. 504 pp (Adv. Mar. Biol.).
De Grave, S" N. D. Pentcheff, S. T. Ahyong, T. Y. Chan, K. A. Crandall, P. C. Dworschak, D. L. Felder, R. M. Feldmann, C. H. J. M. Fransen, L. Y. D. Goulding, R. Lemaitre, M. E. Y. Low, J. W. Martin, P. K. L. Ng, C. E. Schweitzer, S. H. Tan, D. Tshudy & R. Wetzer. 2009. A classification of living and fossil genera of Decapod crustacean. Raffles Bull. Zool. 21:1-109.
de Oliveira, U. O., A. S. da Rosa Araujo, A. Bello-Klein, R. S. da Silva & L. C. Kucharski. 2005. Effects of environmental anoxia and different periods of reoxygenation on oxidative balance in gills of the estuarine crab Chasmagnathus granulala. Comp. Biochem. Physiol. B 140:51-57.
Drose, S. & U. Brandt. 2012. Molecular mechanisms of superoxide production by the mitochondrial respiratory chain. Adv. Exp. Med. Biol. 748:145-169.
Eads, B. D. & S. C. Hand. 2003. Mitochondrial mRNA stability and polyadenylation during anoxia-induced quiescence in the brine shrimp Artemia franciscana. J. Exp. Biol. 206:3681-3692.
El-Khoury, R., E. Dufour, M. Rak. N. Ramanantsoa, N. Grandchamp, Z. Csaba, B. Duvillie. P. Benit, J. Gallego, P. Gressens, C. Sarkis, H. T. Jacobs & P. Rustin. 2013. Alternative oxidase expression in the mouse enables bypassing cytochrome c oxidase blockade and limits mitochondrial ROS overproduction. PLoS Genet. 9:el003182.
Faccenda, D. & M. Campanella. 2012. Molecular regulation of the mitochondrial [F.sub.0][F.sub.1-] ATPsynthase: physiological and pathological significance of the inhibitory factor 1 (IF1). Int. J. Cell Biol. 2012:1-12.
Feng, Y., W. Li, J. Li, J. Wang, J. Ge. D. Xu, Y. Liu, K. Wu, Q. Zeng, J. W. Wu. C. Tian, B. Zhou & M. Yang. 2012. Structural insight into the type-II mitochondrial NADH dehydrogenases. Nature 491:478-482.
Freire, C. A., V. G. Togni & M. Hermes-Lima. 2011. Responses of free radical metabolism to air exposure or salinity stress, in crabs (Callinectes danae and C. ornatus) with different estuarine distributions. Comp. Biochem. Physiol. A 160:291-300.
Fukuda, R., H. Zhang, J. W. Kim, L. Shimoda, C. V. Dang & G. L. Semenza. 2007. HIF-1 regulates cytochrome oxidase subunits to optimize efficiency of respiration in hypoxic cells. Cell 129:111-122.
Ghaffari, N., A. Sanchez-Flores, R. Doan, K. D. Garcia-Orozco, P. L. Chen, A. Ochoa-Leyva, A. A. Lopez-Zavala, J. S. Carrasco. C. Hong, L. G. Brieba, E. Rudino-Pinera, P. D. Blood. J. E. Sawyer, C. D. Johnson, S. V. Dindot, R. R. Sotelo-Mundo & M. F. Criscitiello. 2014. Novel transcriptome assembly and improved annotation of the whiteleg shrimp (Litopenaeus vannamei). a dominant crustacean in global seafood mariculture. Sci. Rep. 4:1-10.
Giorgio. V., S. von Stockum, M. Antoniel, A. Fabbro, F. Fogolari, M. Forte, G. D. Glick, V. Petronilli, M. Zoratti, I. Szabo, G. Lippe & P. Bernardi. 2013. Dimers of mitochondrial ATP synthase form the permeability transition pore. Proc. Natl. Acad. Sci. USA 110:5887-5892.
Glenner, H., P. F. Thomsen, M. B. Hebsgaard, M. V. Sorensen & E. Willerslev. 2006. The origin of insects. Science 314:1883-1884.
Gorr, T. A., J. D. Cahn, H. Yamagata & H. F. Bunn. 2004. Hypoxia-induced synthesis of hemoglobin in the crustacean Daphnia magna is hypoxia-inducible factor-dependent. J. Biol. Chem. 279:36038-36047.
Gorr, T. A., M. Gassmann & P. Wappner. 2006. Sensing and responding to hypoxia via HIF in model invertebrates. J. Insect Physiol. 52:349-364.
Grieshaber, M. K. & S. Volkel. 1998. Animal adaptations for tolerance and exploitation of poisonous sulfide. Annu. Rev. Physiol. 60:33-53.
Guerrero-Castillo, S., D. Araiza-Olivera, A. Cabrera-Orefice, J. Espinasa-Jaramillo, M. Gutierrez-Aguilar, L. A. Luevano-Martinez, A. Zepeda-Bastida & S. Uribe-Carvajal. 2011. Physiological uncoupling of mitochondrial oxidative phosphorylation. Studies in different yeasts species. J. Bioenerg. Biomembr. 43:323-331.
Guzy, R. D. & P. T. Schumacker. 2006. Oxygen sensing by mitochondria at complex III: the paradox of increased reactive oxygen species during hypoxia. Exp. Physiol. 91:807-819.
Halestrap, A. P. 2009. What is the mitochondrial permeability transition pore? J. Mol. Cell. Cardiol. 46:821-831.
Halestrap, A. P. & A. M. Davidson. 1990. Inhibition of [Ca.sup.2+] -induced large-amplitude swelling of liver and heart mitochondria by cyclosporin is probably caused by the inhibitor binding to mitochondrial-matrix peptidyl-prolyl cis-trans isomerase and preventing it interacting with the adenine nucleotide translocase. Biochem. J. 268:153-160.
Halestrap, A. P., P. M. Kerr, S. Javadov & K. Y. Woodfield. 1998. Elucidating the molecular mechanism of the permeability transition pore and its role in reperfusion injury of the heart. Biochim. Biophys. Acta 1366:79-94.
Hardy, K. M., C. R. Follett, L. E. Burnett & S. C. Lema. 2012. Gene transcripts encoding hypoxia-inducible factor (HIF) exhibit tissue-and muscle fiber type-dependent responses to hypoxia and hyper-capnic hypoxia in the Atlantic blue crab. Callinectes sapidus. Comp. Biochem. Physiol. A 163:137-146.
Hermes-Lima, M. & T. Zenteno-Savin. 2002. Animal response to drastic changes in oxygen availability and physiological oxidative stress. Comp. Biochem. Physiol. C Toxicol. Pharmacol. 133:537-556.
Hildebrandt, T. M. & M. K. Grieshaber. 2008. Redox regulation of mitochondrial sulfide oxidation in the lugworm, Arenicola marina. J. Exp. Biol. 211:2617-2623.
Hinkle, P. C. 2005. P/O ratios of mitochondrial oxidative phosphorylation. Biochim. Biophys. Acta 1706:1-11.
Hochachka, P. W. & P. L. Lutz. 2001. Mechanism, origin, and evolution of anoxia tolerance in animals. Comp. Biochem. Physiol. B 130:435-459.
Holman. J. D. & S. C. Hand. 2009. Metabolic depression is delayed and mitochondrial impairment averted during prolonged anoxia in the ghost shrimp, Lepidophthalmus louisianensis (Schmitt, 1935). J. Exp. Mar. Biol. Ecol. 376:85-93.
Iwata, M., Y. Lee, T. Yamashita, T. Yagi, S. Iwata, A. D. Cameron & M. J. Maher. 2012. The structure of the yeast NADH dehydrogenase (Ndil) reveals overlapping binding sites for water- and lipid-soluble substrates. Proc. Natl. Acad. Sci. USA 109:15247-15252.
Ji, Ch., G. Weiran, M. Zhang, X. Lu. Y. Ni & X. Guo. 2012. Caenorhabditis elegans ucp-4 regulates fat metabolism: supression of ucp-4 expression induced obese phenotype and caused impairment of insulin like pathway. Gene 491:158-164.
Jimenez-Gutierrez. L. R., J. Hernandez-Lopez, M. A. Islas-Osuna & A. Muhlia-Almazan. 2013. Three nucleus-encoded subunits of mitochondrial cytochrome c oxidase of the whiteleg shrimp Litopenaeus vannamei: cDNA characterization, phylogeny and mRNA expression during hypoxia and reoxygenation. Comp. Biochem. Physiol. B 166:30-39.
Jimenez-Gutierrez, L. R., S. Uribe-Carvajal, A. Sanchez-Paz, C. Chimeo & A. Muhlia-Almazan. 2014. The cytochrome c oxidase and its mitochondrial function in the whiteleg shrimp Litopenaeus vannamei during hypoxia. J. Bioenerg. Biomembr. 46:189-196.
Johns, A. R., A. C. Taylor, R. J. A. Atkinson & M. K. Grieshaber. 1997. Sulphide metabolism in thalassinidean Crustacea. J. Mar. Biol. Assoc. U.K. 77:127-144.
Johnson, J. G., M. R. Paul, C. D. Kniffin, P. E. Anderson, L. E. Burnett & K. G. Burnett. 2015. High C[O.sub.2] alters the hypoxia response of the Pacific whiteleg shrimp (Litopenaeus vannamei) transcriptome including known and novel hemocyanin isoforms. Physiol. Genomics 47:548-558.
Kadenbach, B. 2003. Intrinsic and extrinsic uncoupling of oxidative phosphorylation. Biochim. Biophys. Acta 1604:77-94.
Kern, B., A. V. Ivanina, H. Piontkivska, E. P. Sokolov & I. M. Sokolova. 2009. Molecular characterization and expression of a novel homolog of uncoupling protein 5 (UCP5) from the eastern oyster Crassostrea virginica (Bivalvia: Ostreidae). Comp. Biochem. Physiol. Part D Genomics Proteomics 4:121-127,
Kilpert, F. & L. Podsiadlowski. 2006. The complete mitochondrial genome of the common sea slater, Ligia oceanica (Crustacea, Isopoda) bears a novel gene order and unusual control region features. BMC Genomics 7:1-18.
Konrad. C, G. Kiss. B. Torocsik, V. Adam-Vizi & C. Chinopoulos. 2012. Absence of Ca" (+)-induced mitochondrial permeability transition but presence of bongkrekate-sensitive nucleotide exchange in C. crangon and P. serratus. PLoS One 7:e39839.
Kowaltowski, A. J., N. C. de Souza-Pinto. R. F. Castilho & A. E. Vercesi. 2009. Mitochondria and reactive oxygen species. Free Radic. Biol. Med. 47:333-343.
Kroemer, G., L. Galluzi & C. Brenner. 2007. Mitochondrial membrane permeabilization in cell death. Physiol. Rev. 87:99-163.
Krumschnabel, G., C. Manzl, C. Berger & B. Hofer. 2005. Oxidative stress, mitochondrial permeability transition, and cell death in Cu-exposed trout hepatocytes. Toxicol. Appl. Pharmacol. 209:62-73.
Kwast. K. E. & S. C. Hand. 1996. Acute depression of mitochondrial protein synthesis during anoxia: contributions of oxygen sensing, matrix acidification, and redox state. J. Biol. Chem. 271:7313-7319.
Lafferty. K. D. & A. M. Kuris. 2009. Parasitic castration: the evolution and ecology of body snatchers. Trends Parasitol. 25:564-572.
Lane, N. 2002. Oxygen, the molecule that made the world. USA: Oxford University Press. OUP Oxford Publisher. 374 pp.
Lawniczak, M., C. Romestaing, D. Roussel. C. Maazousi, D. Renault & F. Hervant. 2013. Preventive antioxidant responses to extreme oxygen level fluctuation in a subterranean crustacean. Comp. Biochem. Physiol. A 165:299-303.
Lesser, M. P. 2006. Oxidative stress in marine environments: biochemistry and physiological ecology. Anmt. Rev. Physiol. 68:253-278.
Li, T. & M. Brouwer. 2007. Hypoxia-inducible factor, gsHIF, of the grass shrimp Palaemonetes pugio: molecular characterization and response to hypoxia. Comp. Biochem. Physiol. B 147:11-19.
Lutz, P. L. & S. L. Milton. 2004. Negotiating brain anoxia survival in the turtle. J. Exp. Biol. 207:3141-3147.
Lutz. P. L. & G. E. Nilsson. 1997. Contrasting strategies for anoxic brain survival-glycolysis up or down. J. Exp. Biol. 200:411-419.
Machida, R. J., M. U. Miya, M. Nishida & S. Nishida. 2002. Complete mitochondrial DNA sequence of Tigriopus japonicus (Crustacea: Copepoda). Mar. Bioteclmol. (NY) 4:406-417.
Martinez-Cruz, O., A. Arvizu-Flores, R. R. Sotelo-Mundo & A. Muhlia-Almazan. 2015. The nuclear encoded subunits gamma, delta and epsilon from the shrimp mitochondrial [F.sub.1] -ATP synthase, and their transcriptional response during hypoxia. J. Bioenerg. Biomembr. 47:223-234.
Martinez-Cruz. O., A. M. Calderon de la Barca, S. Uribe-Carvajal & A. Muhlia-Almazan. 2012. The function of mitochondrial [F.sub.0][F.sub.1] ATP-synthase from the whiteleg shrimp Litopenaeus vannamei muscle during hypoxia. Comp. Biochem. Physiol. B 162:107- 112.
Martinez-Cruz, O., F. Garcia-Carreno, A. Robles-Romo, A. Varela-Romero & A. Muhlia-Almazan. 2011. Catalytic subunits atpa and atpfi from the Pacific white shrimp Litopenaeus vannamei [F.sub.0][F.sub.1], ATP-synthase complex: cDNA sequences, phylogenies, and mRNA quantification during hypoxia. J. Bioenerg. Biomembr. 43:119-133.
McDonald, A. E., G. C. Vanlerberghe & J. F. Staples. 2009. Alternative oxidase in animals: unique characteristics and taxonomic distribution. J. Exp. Biol. 212:2627-2634.
Mendez-Romero, O. A. 2014. Caracterizacion del ADN complementario de las subunidades nucleares de la citocromo c oxidase COX Via. COX VIb y COX Vic del camaron bianco del Pacifico Litopenaeus vannamei. Bachelor thesis, Instituto Tecnologico de Sonora, Sonora, Mexico.
Mendez-Romero, O. A. 2016. Las proteinas mitocondriales desacoplantes UCP4 y UCP5 del camaron bianco y su posible funcion durante la hipoxia. MSc thesis, Centro de Investigacion en Alimentacion y Desarrollo, A.C., Sonora, Mexico.
Menze, M. A., G. Fortner, S. Nag & S. C. Hand. 2010. Mechanisms of apoptosis in Crustacea: what conditions induce versus suppress cell death? Apoptosis 15:293-312.
Menze. M. A., K. Hutchinson. S. M. Laborde & S. C. Hand. 2005. Mitochondrial permeability transition in the crustacean Artemia franciscana: absence of a calcium-regulated pore in the face of profound calcium storage. Am. J. Physiol. Regul. Integr. Comp. Physiol. 289:R68-R76.
Minchenko, O., I. Opentanova & J. Caro. 2003. Hypoxic regulation of the 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase gene family (PFKFB-1-4) expression in vivo. FEBS Lett. 554:264-270.
Mitchell, P. 1966. Chemiosmotic coupling in oxidative and photosynthetic phosphorylation. Biol. Rev. Camb. Philos. Soc. 41:445-502.
Miwa, S., J. St-Pierre, L. Partridge & M. D. Brand. 2003. Superoxide and hydrogen peroxide production by Drosophila mitochondria. Free Radic. Biol. Med. 35:938-948.
Mracek. T., Z. Drahota & J. Houstek. 2013. The function and the role of the mitochondrial glycerol-3-phosphate dehydrogenase in mammalian tissues. Biochim. Biophys. Acta 1827:401-410.
Muhlia-Almazan, A., O. Martinez-Cruz, M. A. Navarrete del Toro, F. Garcia-Carrefio, R. Arreola. R. Sotelo-Mundo & G. Yepiz-Plascencia. 2008. Nuclear and mitochondrial subunits from the white shrimp Litopenaeus vannamei [F.sub.0][F.sub.1] ATP-synthase complex: cDNA sequence, molecular modeling, and mRNA quantification of atp9 and atp6. J. Bioenerg. Biomembr. 40:359-369.
Murphy. M. P. 2009. How mitochondria produce reactive oxygen species. Biochem. J. 417:1-13.
Nilsson, G. E. & P. L. Lutz. 2004. Anoxia tolerant brains. J. Cereb. Blood Flow Metab. 24:475-486.
O'Brien, J. & R. D. Vetter. 1990. Production of thiosulphate during sulphide oxidation by mitochondria of the symbiont-containing bivalve Solemya reidi. J. Exp. Biol. 149:133-148.
Parrilla-Taylor, D. P. & T. Zenteno-Savin. 2011. Antioxidant enzyme activities in Pacific white shrimp (Litopenaeus vannamei) in response to environmental hypoxia and reoxygenation. Aquaculture 318:379-383.
Poltermann, M., H. Hop & S. Falk-Petersen. 2000. Life under Arctic sea ice--reproduction strategies of two sympagic (ice-associated) amphipod species, Gammarus wilkitzkii and Apherusa glacialis. Mar. Biol. 136:913-920.
Powell, M. A. & G. N. Somero. 1986. Hydrogen sulfide oxidation is coupled to oxidative phosphorylation in mitochondria of Solemya reidi. Science 233:563-566.
Puente-Carreon, E. 2009. Respuestas fisiologicas de juveniles de camaron bianco Litopenaeus vannamei, a condiciones oscilantes de oxigeno disuelto y temperatura. PhD thesis, Instituto Politecnico Nacional, Baja California Sur, Mexico.
Regier, J. C, J. W. Shultz, A. Zwick, A. Hussey, B. Ball, R. Wetzer, J. W. Martin & C. W. Cunningham. 2010. Arthropod relationships revealed by phylogenomic analysis of nuclear protein-coding sequences. Nature 463:1079-1083.
Rehm, P., J. Borner, K. Meusemann, B. M. von Reumont, S. Simon, H. Hadrys, B. Misof & T. Burmester. 2011. Dating the arthropod tree based on large-scale transcriptome data. Mol. Phylogenet. Evol. 61:880-887.
Romero, M. C, M. Ansaldo & G. A. Lovrich. 2007. Effect of aerial exposure on the antioxidant status in the subantarctic stone crab Paralomis granulosa (Decapoda:Anomura). Comp. Biochem. Physiol. C Toxicol. Pharmacol. 146:54-59.
Sanderson, T. H., C. A. Reynolds, R. Kumar, K. Pfzyklenk & M. Huttemann. 2013. Molecular mechanisms of ischemia-reperfusion injury in brain: pivotal role of the mitochondrial membrane potential in reactive oxygen species generation. Mol. Neurobiol. 47:9-23.
Segawa, R. D. & T. Aotsuka. 2005. The mitochondrial genome of the Japanese freshwater crab, Geothelphusa dehaani (Crustacea: Brachyura): evidence for its evolution via gene duplication. Gene 355:28-39.
Semenza, G. L. 2000. HIF-1: mediator of physiological and pathophysiological responses to hypoxia. J. Appl. Physiol. 88:1474-1480.
Shen, H., Y. Hu, Y. Ma, X. Zhou, Y. Shui, C. Li, P. Xu & X. Sun. 2014. In-depth transcriptome analysis of the red swamp crayfish Procambarus clarkii. PLoS One 9:el 10548.
Shen, X., J. Ren, Z. Cui, Z. Sha, B. Wang, J. Xiang & B. Liu. 2007. The complete mitochondrial genomes of two common shrimps (Litopenaeus vannamei and Fenneropenaeus chinensis) and their phylogenomic considerations. Gene 403:98-109.
Shen, X., M. Tian, B. Yan & K. Chu. 2015. Phylomitogenomics of Malacoslraca (Arthropoda: Crustacea). Acta Oceanol. Sin. 34:84-92.
Skodova, I., Z. Verner, F. Bringaud, P. Fabian, J. Lukes & A. Horvath. 2013. Characterization of two mitochondrial flavin adenine dinucleotide-dependent glycerol-3-phosphate dehydrogenases in Trypanosoma brucei. Eukaryot. Cell 12:1664-1673.
Slocinska, M., N. Antos-Krzeminska, G. Rosinski & W. Jarmuszkiewicz. 2011. Identification and characterization of uncoupling protein 4 in fat body and muscle mitochondria from the cockroach Gromphadorhina cocquereliana. J. Bioenerg. Biomembr. 43:717-727.
Smit, N. J., N. L. Bruce & K. A. Hadfield. 2014. Global diversity offish parasitic isopod crustaceans of the family Cymothoidae. Int. J. Parasitol. Parasites Wildl. 3:188-197.
Sokolova, I. M., S. Evans & F. M. Hughes. 2004. Cadmium-induced apoptosis in oyster hemocytes involves disturbance of cellular energy balance but no mitochondrial permeability transition. J. Exp. Biol. 207:3369-3380.
Sokolova, I. M. & E. P. Sokolov. 2005. Evolution of mitochondrial uncoupling proteins: novel invertebrate UCP homologues suggest early evolutionary divergence of the UCP family. FEBS Lett. 579:313-317
Solaini, G. & D. A. Harris. 2005. Biochemical dysfunction on heart mitochondria exposed to ischemia and reperfusion. Biochem. J. 390:377-394.
Sonanez-Organis, J. G., A. B. Peregrino-Uriate, S. Gomez-Jimenez, A. Lopez-Zavala, H. J. Forman & G. Yepiz-Plasencia. 2009. Molecular characterization of hypoxia inducible factor-1 (HIF-1) from the white shrimp Litopenaeus vannamei and tissue-specific expression under hypoxia. Comp. Biochem. Physiol. C Toxicol. Pharmacol. 150:395-405.
Staton, J. L., L. L. Daehler & W. M. Brown. 1997. Mitochondrial gene arrangement of the horseshoe crab Limulus polyphemus L.: conservation of major features among arthropod classes. Mol. Biol. Evol. 14:867-874.
Storey, K. B. 2004. Adventures in oxygen metabolism. Comp. Biochem. Physiol. B 139:359-369.
Sun, S., F. Xuan, H. Fu, J. Zhu, X. Ge & Z. Gu. 2015. Transcriptomic and histological analysis of hepatopancreas, muscle and gill tissues of oriental river prawn (Macrobrachium nipponense) in response to chronic hypoxia. BMC Genomics 16:1-13.
Sussarellu, R., C. Fabioux, M. Camacho Sanchez, N. Le Goic, C. Lambert, P. Soudant & D. Moraga. 2012. Molecular and cellular response to short-term oxygen variations in the Pacific oyster Crassostrea gigas. J. Exp. Mar. Biol. Ecol. 412:87-95.
Theissen, U. & W. Martin. 2008. Sulfide:quinone oxidoreductase (SQR) from the lugworm Arenicola marina shows cyanide- and thioredoxin-dependent activity. FEBS J. 275:1131-1139.
Valverde, J. R., B. Batuecas, C. Moratilla, R. Marco & R. Garesse. 1994. The complete mitochondrial DNA sequence of the crustacean Artemia franciscana. J. Mol. Evol. 39:400-408.
Vaquer-Sunyer, R. & C. M. Duarte. 2008. Thresholds of hypoxia for marine biodiversity. Proc. Natl. Acad. Sci. USA 105:15452-15457.
von Stockum, S., V. Giorgio, E. Trevisan, G. Lippe. G. D. Glick, M. A. Forte, C. Da-Re, V. Checchetto, G. Mazzotta, R. Costa, I. Szabo & P. Bernardi. 2015. F-ATPase of Drosophila melanogaster forms 53-Picosiemen (53-Ps) channels responsible for mitochondrial [Ca.sup.2+]-induced [Ca.sup.2+] release. J. Biol. Chem. 290:4537-4544.
Vrbacky, M., Z. Drahota, T. Mracek, A. Vojtiskova, P. Jesina, P. Stopka & J. Houstek. 2007. Respiratory chain components involved in the glycerophosphate dehydrogenase-dependent ROS production by brown adipose tissue mitochondria. Biochim. Biophys. Acta 1767:989-997.
Welker, A. F., D. C. Moreira, E. G. Campos & M. Hermes-Lima. 2013. Role of redox metabolism for adaptation of aquatic animals to drastic changes in oxygen availability. Comp. Biochem. Physiol. A 165:384-404.
Werner, I. & H. Auel. 2005. Seasonal variability in abundance, respiration and lipid composition of Artie under-ice amphipods. Mar. Ecol. Prog. Ser. 292:251-262.
Wilson, K., V. Cahill, E. Bailment & J. Benzie. 2000. The complete sequence of the mitochondrial genome of the crustacean Penaeus monodon: are malacostracan crustaceans more closely related to insects than to branchiopods? Mol. Biol. Evol. 17:863-874.
Wu, R. S., P. K. Lam & K. L. Wan. 2002. Tolerance to, and avoidance of, hypoxia by the penaeid shrimp (Metapenaeus ensis). Environ. Pollut. 118:351-355.
Yamauchi, M. M., M. U. Miya & M. Nishida. 2003. Complete mitochondrial DNA sequence of the swimming crab, Portunus trituberculalus (Crustacea: Decapoda: Brachyura). Gene 311:129-135.
Zenteno-Savin, T., R. Saldierna & M. Ahuejote-Sandoval. 2006. Superoxide radical production in response to environmental hypoxia in cultured shrimp. Comp. Biochem. Physiol. C Toxicol. Pharmacol. 142:301-308.
Zilli, L., R. Schiavone, G. Scordella, V. Zonno, T. Verri, C. Storelli & S. Vilella. 2003. Changes in cell type composition and enzymatic activities in the hepatopancreas of Marsupenaeus japonicus during the moulting cycle. J. Comp. Physiol. B 173:355-363.
Zilli, L., R. Schiavone, C. Storelli & S. Vilella. 2007. Analysis of calcium concentration fluctuations in hepatopancreatic R cells of Marsupenaeus japonicas during the molting cycle. Biol. Bull. 212:161-168.
OLIVIERT MARTINEZ-CRUZ, (1) CINDY CHIMEO, (2) CHRYSTIAN M. RODRIGUEZ-ARMENTA (2) AND ADRIANA MUHLIA-ALMAZAN (2*)
(1) Departamento de Investigation y Posgrado en Alimentos, Universidud de Sonora, Boulevard Luis Encinas SjN. Col. Centra, Hermosillo, Sonora 83000, Mexico; 'Bioenergetics and Molecular Genetics Lab, Centro de Investigation en Alimentation y Desarrollo A.C. Carretera a la Victoria km 0.6, Hermosillo, Sonora 1735, Mexico
(*)Corresponding author. E-mail: email@example.com
TABLE 1. The deduced proteins of nuclear-encoded transcripts involved in the mitochondrial function of the white shrimp Litopenaeus vannamei. Similar to other Crustaceans transcripts or expressed sequence tags Deduced protein name from (GenBank access no.) L. vannamei transcripts Electron transport chain complex I NADH dehydrogenase subunit F L. vannamei FEI01646.1 - NADH dehydrogenase Penaeus monodon [ubiquinone] flavoprotein 2 GE615804.1 - NADH dehydrogenase L. vannamei [ubiquinone] flavoprotein 3 FE 126620.1 - NADH dehydrogenase L. vannamei [ubiquinone] 1 beta subcomplex subunit 4 FE053783.1 - NADH dehydrogenase No record [ubiquinone] 1 alpha subcomplex subunit 5 - - NADH dehydrogenase No record [ubiquinone] 1 alpha subcomplex subunit 6 - - NADH dehydrogenase P. monodon [ubiquinone] iron-sulfur protein 7 GW995271.1 - NADH dehydrogenase P. monodon [ubiquinone] 1 alpha subcomplex subunit 8 EE662014.1 - NADH dehydrogenase L. vannamei [ubiquinone] 1 alpha subcomplex subunit 9 FE185623.1 - NADH dehydrogenase L. vannamei [ubiquinone] 1 alpha subcomplex subunit 11 FE156411.1 - NADH dehydrogenase No record [ubiquinone] 1 alpha subcomplex subunit 12 - - Electron transport chain complex II Succinate dehydrogenase subunit A L. vannamei FE050092.1 - Succinate dehydrogenase subunit B L. vannamei FE103086.1 - Electron transport chain complex III Cytochrome b5 L. vannamei FE126775.1 - Cytochrome b-cl complex subunit Rieske L. vannamei FE 154972.1 - Cytochrome b-cl complex subunit 1 L. vannamei FE103284.1 - Cytochrome b-cl complex subunit 2 P. monodon GW995086.1 - Cytochrome b-cl complex subunit 7 L. vannamei FE150813.1 - Cytochrome cl P. monodon GO073581.1 - Electron transport chain complex IV Cytochrome c L. vannamei FE126543.1 - COX 7 subunit A L. vannamei FE153349.1 - COX 11 L. vannamei FE075323.1 - COX 15 P. monodon GE615894.1 - COX 16 P. monodon GW996288.1 Electron transport chain complex V ATP synthase subunit D L. vannamei FE123815.1 - ATP synthase subunit E L. vannamei GR973479.1 - ATP synthase subunit F L. vannamei FE137182.1 - ATP synthase subunit G Fenneropenaeus indicus GT968440.1 - ATP synthase subunit O L. vannamei FE157827.1 - Krebs cycle Citrate synthase L. vannamei FE103408.1 Aconitase L. vannamei FE135452.1 - Succinyl-CoA synthetase L. vannamei subunit alpha FE076651.1 - ADP-forming Succinyl-CoA L. vannamei ligase subunit beta FE051092.1 - Mitochondrial NADP(+)-dependent L. vannamei Isocitrate dehydrogenase FE107598.1 - Mitochondrial NAD(+)-Isocitrate L. vannamei dehydrogenase subunit alpha CK572301.1 - Mitochondrial NAD(+)-Isocitrate L. vannamei dehydrogenase subunit beta FE061769.1 - Mitochondrial NAD(+)-Isocitrate L. vannamei dehydrogenase subunit gamma FE 174200.1 - Malate dehydrogenase L. vannamei FE123204.1 - Fumarase No record - - 2-Oxoglutarate dehydrogenase L. vannamei FE178128.1 - Dihydrolipoyllysine-residue P. monodon succinyltransferase GE615760.1 - Pyruvate dehydrogenase complex Pyruvate dehydrogenase El L. vannamei component subunit alpha FE085445.1 - Pyruvate dehydrogenase El Metapenaeus ensis component subunit beta CV179091.1 - Dihydrolipoyllysine-residue P. monodon acetyltransferase GE616028.1 - Dihydrolipoyl dehydrogenase P. monodon GE615629.1 - Mitochondrial carriers, permeability transition, and calcium transport proteins Voltage-dependent anion channel L. vannamei FE102731.1 - Adenine nucleotide P. monodon translocator isoform 3 GW995486.1 - Cyclophilin D L. vannamei FE087681.1 - Phosphate carrier protein L. vannamei FE149241.1 - Similar to other Deduced protein name from species proteins: L. vannamei transcripts Electron transport chain complex I NADH dehydrogenase subunit F Aplysia californica Komagataeibacter intermedins TF2 Rhizopus delemar RA 99-880 NADH dehydrogenase Paramecium tetraurelia [ubiquinone] flavoprotein 2 strain d4-2 Tetrahymena thermophila SB2I0 Salpingoeca rosetta NADH dehydrogenase Acyrthosiphon pisum [ubiquinone] flavoprotein 3 Peromyscus maniculatus hairdii Rattus norvegicus NADH dehydrogenase Zootermopsis nevadensis [ubiquinone] 1 beta subcomplex subunit 4 Diaphorina cilri Sparus aurata NADH dehydrogenase Camelina saliva [ubiquinone] 1 alpha subcomplex subunit 5 Zootermopsis nevadensis Brassica rapa NADH dehydrogenase Phoenix dactylifera [ubiquinone] 1 alpha subcomplex subunit 6 Jatropha curcas Brassica rapa NADH dehydrogenase Zootermopsis nevadensis [ubiquinone] iron-sulfur protein 7 Plutella xylostella Danaus plexippus NADH dehydrogenase Zootermopsis nevadensis [ubiquinone] 1 alpha subcomplex subunit 8 Bactrocera dorsalis Drosophila melanogaster NADH dehydrogenase Ichthyophthirius [ubiquinone] 1 alpha subcomplex multifiliis subunit 9 Solanum lycopersicum Tarenaya hassleriana NADH dehydrogenase Bactrocera cucurbitae [ubiquinone] 1 alpha subcomplex subunit 11 Ceratitis capitata Musca domestica NADH dehydrogenase Tetrahymena thermophila [ubiquinone] 1 alpha subcomplex SB210 subunit 12 Ichthyophthirius multifiliis Stylonychia lemnae Electron transport chain complex II Succinate dehydrogenase subunit A Paramecium tetraurelia strain d4-2 Tetrahymena thermophila SB210 Moesziomyces antarclicus T-34 Succinate dehydrogenase subunit B Zootermopsis nevadensis Lysiphlebus testaceipes Anopheles darlingi Electron transport chain complex III Cytochrome b5 Tribolium castaneum Zootermopsis nevadensis Anopheles darlingi Cytochrome b-cl complex subunit Rieske Tribolium castaneum Culex quinquefasciatus Oreochromis niloticus Cytochrome b-cl complex subunit 1 Homo sapiens Pongo abelii Tarsius syrichta Cytochrome b-cl complex subunit 2 Tribolium castaneum Acromyrmex echinatior Salmo salar Cytochrome b-cl complex subunit 7 Musca domestica Ceratitis capitata Bactrocera dorsalis Cytochrome cl Tetrahymena thermophila Stylonychia lemnae Oxytricha trifallax Electron transport chain complex IV Cytochrome c Marsupenaeus japonicus Tigriopus californicus Locusta migratoria COX 7 subunit A Zootermopsis nevadensis Musca domestica Bactrocera cucurbitae COX 11 Bactrocera cucurbitae Ceratitis capitata Culex quinquefasciatus COX 15 Culex quinquefasciatus Anopheles darlingi Danio rerio COX 16 Tribolium castaneum Zootermopsis nevadensis Bombyx mori Electron transport chain complex V ATP synthase subunit D Solenopsis invicta Bombus terreslris Anopheles darlingi ATP synthase subunit E Aedes albopictus Apis mellijera Ceratitis capilata ATP synthase subunit F Anopheles darlingi Culex quinquefasciatus Aedes aegypti ATP synthase subunit G Palaemon varians Ixodes scapularis Musca domestica ATP synthase subunit O Coptotermes formosanus Linepithema humile Tribolium caslaneum Krebs cycle Citrate synthase Anopheles darlingi Sus scrofa Danio rerio Aconitase Lasius niger Daphnia magna Daphnia pulex Succinyl-CoA synthetase Anopheles darlingi subunit alpha Culex quinquefasciatus Zootermopsis nevadensis ADP-forming Succinyl-CoA Riptortus pedestris ligase subunit beta Zootermopsis nevadensis Tribolium castaneum Mitochondrial NADP(+)-dependent Zootermopsis nevadensis Isocitrate dehydrogenase Salmo salar Astyanax mexicanus Mitochondrial NAD(+)-Isocitrate Crassostrea gigas dehydrogenase subunit alpha Strongyloides ratti Aplysia californica Mitochondrial NAD(+)-Isocitrate Musca domestica dehydrogenase subunit beta Bombyx mori Linepithema humile Mitochondrial NAD(+)-Isocitrate Tribolium caslaneum dehydrogenase subunit gamma Diaphorina citri Harpegnathos saltator Malate dehydrogenase Ictalurus pune talus Danio rerio Daphnia pulex Fumarase Tribolium castaneum Zootermopsis nevadensis Danio rerio 2-Oxoglutarate dehydrogenase Apis mellifera Bomhus terrestres Tribolium castaneum Dihydrolipoyllysine-residue Culex quinquefasciatus succinyltransferase Bombyx mori Musca domestica Pyruvate dehydrogenase complex Pyruvate dehydrogenase El Tetrahymena thermophila component subunit alpha SB210 Oxytricha trifallax Euplotes sp. BB-2004 Pyruvate dehydrogenase El Zootermopsis nevadensis component subunit beta Danio rerio Apis mellifera Dihydrolipoyllysine-residue Crassostrea gigas acetyltransferase Lingula anatina Biomphalaria glabrata Dihydrolipoyl dehydrogenase Zootermopsis nevadensis Rhy-opertha dominica Tribolium castaneum Mitochondrial carriers, permeability transition, and calcium transport proteins Voltage-dependent anion channel Eriocheir sinensis Culex quinquefasciatus Anopheles darlingi Adenine nucleotide Poecilia Formosa translocator isoform 3 Stegastes partitus Cynoglossus semilaevis Cyclophilin D Poecilia Formosa Larimichthys crocea Danio rerio Phosphate carrier protein Lingula anatina Stegodyphus mimosarum Crassostrea gigas GenBank Deduced protein name from access no. Total L. vannamei transcripts score Electron transport chain complex I NADH dehydrogenase subunit F XP_005097475.1 170 GAN86293.1 170 EIE75829.1 170 NADH dehydrogenase XP 001347081.1 298 [ubiquinone] flavoprotein 2 XP_001017531.1 279 XP_004990429.1 226 NADH dehydrogenase XP_003240259.1 47.4 [ubiquinone] flavoprotein 3 XP_006973964.1 43.5 NP_072129.2 40.8 NADH dehydrogenase KDR12405.1 138 [ubiquinone] 1 beta subcomplex subunit 4 XP_008477125.1 112 AGV76785.1 77.4 NADH dehydrogenase XP_010442830.1 50.4 [ubiquinone] 1 alpha subcomplex subunit 5 KDR17347.1 45.1 XP_009119817.1 49.7 NADH dehydrogenase XP_008775040.1 57.8 [ubiquinone] 1 alpha subcomplex subunit 6 XP_012068683.1 57.4 XP 009120779.1 55.1 NADH dehydrogenase KDR 14423.1 300 [ubiquinone] iron-sulfur protein 7 XP_011549720.1 292 EHJ72892.1 288 NADH dehydrogenase KDR15899.1 223 [ubiquinone] 1 alpha subcomplex subunit 8 XP_011202093.1 214 NP_611954.2 207 NADH dehydrogenase XP_004034899.1 280 [ubiquinone] 1 alpha subcomplex subunit 9 XP_004247128.1 169 XP_010548735.1 169 NADH dehydrogenase XP_011196653.1 129 [ubiquinone] 1 alpha subcomplex subunit 11 XP_004522669.1 124 XP_005179354.1 98.6 NADH dehydrogenase XP_001471372.1 152 [ubiquinone] 1 alpha subcomplex subunit 12 XP_004027195.1 138 CDW76573.1 127 Electron transport chain complex II Succinate dehydrogenase subunit A XP_001347005.1 452 XP_001014703.2 447 GAC72599.1 444 Succinate dehydrogenase subunit B KDR22350.1 449 AAY63983.1 435 ETN61683.1 424 Electron transport chain complex III Cytochrome b5 XP_975884.1 369 KDR21019.1 352 ETN57924.1 342 Cytochrome b-cl complex subunit Rieske NP_001164310.1 328 XP_001867379.1 322 XP_003447114.1 319 Cytochrome b-cl complex subunit 1 NP_003356.2 99.8 XP_009237316.1 99.8 XP_008066364.1 98.2 Cytochrome b-cl complex subunit 2 XP_975769.1 315 XP_011059867.1 302 ACN 10092.1 287 Cytochrome b-cl complex subunit 7 XP_005181276.1 120 XP_004529599.1 117 XP_011198166.1 115 Cytochrome cl XP_001027842.2 407 CDW90024.1 270 EJY73104.1 266 Electron transport chain complex IV Cytochrome c BAJ22990.1 202 AAC80550.1 186 AGF80276.1 185 COX 7 subunit A K.DR 19493.1 70.1 XP_005188857.1 62.8 XP_011178340.1 59.7 COX 11 XP_011188067.1 326 XP_004522467.1 320 XP_001865568.1 311 COX 15 XP_001868025.1 466 ETN60022.1 463 AAH66452.1 409 COX 16 XP_972022.1 97.1 KDR21754.1 84.7 XP_004929682.1 80.1 Electron transport chain complex V ATP synthase subunit D XP_011161901.1 352 XP_003397933.1 343 ETN66668.1 335 ATP synthase subunit E AAV90734.1 82.4 XP_624249.1 79.7 XP_004524140.1 73.6 ATP synthase subunit F ETN65643.1 170 XP_001846737.1 167 ABF18130.1 164 ATP synthase subunit G ACR54103.1 173 AAY66986.1 127 XP_005182908.1 119 ATP synthase subunit O AGM32184.1 248 XP_012220907.1 248 XP_968733.1 244 Krebs cycle Citrate synthase ETN60073.1 753 NP_999441.1 726 AAI66040.1 719 Aconitase KMQ95155.1 1297 KZS13121.1 1266 CAB72317.1 1259 Succinyl-CoA synthetase ETN59000.1 455 subunit alpha XP_001868803.1 431 KDR08649.1 459 ADP-forming Succinyl-CoA BAN21224.1 595 ligase subunit beta KDR17135.1 583 XP_970725.1 583 Mitochondrial NADP(+)-dependent KDR14081.1 687 Isocitrate dehydrogenase NP_001133196.1 642 XP_007254547.1 644 Mitochondrial NAD(+)-Isocitrate XP_011425087.1 546 dehydrogenase subunit alpha CEF67094.1 546 XP_005092712.1 543 Mitochondrial NAD(+)-Isocitrate XP_005184353.1 467 dehydrogenase subunit beta XPJH2546738.1 462 XP_012234405.1 472 Mitochondrial NAD(+)-Isocitrate XP_008193347.1 479 dehydrogenase subunit gamma XP_008479949.1 474 XP_011147544.1 446 Malate dehydrogenase NP_001188130.1 531 NP_998296.1 526 EFX69032.1 516 Fumarase XP_967085.1 793 KDR21372.1 765 NP_957257.1 758 2-Oxoglutarate dehydrogenase XP_006566130.1 1510 XP_012168263.1 1499 XP_008193113.1 1505 Dihydrolipoyllysine-residue XP_001845679.1 511 succinyltransferase XP_012546089.1 494 XP_005183707.1 501 Pyruvate dehydrogenase complex Pyruvate dehydrogenase El XP_001017076.2 338 component subunit alpha EJY81740.1 348 AAV32066.1 327 Pyruvate dehydrogenase El KDR20584.1 548 component subunit beta NP_998319.1 503 NP_001229442.1 532 Dihydrolipoyllysine-residue XP_011412451.1 171 acetyltransferase XP_013383918.1 181 XP_013084075.1 177 Dihydrolipoyl dehydrogenase KDR 16622.1 781 AFP20522.1 776 NP_001280524.1 770 Mitochondrial carriers, permeability transition, and calcium transport proteins Voltage-dependent anion channel ADJ94951.2 541 XP_00184263 7.1 380 ETN58314.1 373 Adenine nucleotide XP_007557479.1 344 translocator isoform 3 XP_008274460.1 339 XP_008 324945.1 340 Cyclophilin D XP_007551094.1 390 XP_010746147.1 388 NP_001002065.1 384 Phosphate carrier protein XP_013385882.1 488 KFM69008.1 488 XP_011432203.1 499 Deduced protein name from Query cover (%) % Ident L. vannamei transcripts Electron transport chain complex I NADH dehydrogenase subunit F 98 75 99 74 98 74 NADH dehydrogenase 87 65 [ubiquinone] flavoprotein 2 89 61 85 53 NADH dehydrogenase 18 57 [ubiquinone] flavoprotein 3 25 41 20 39 NADH dehydrogenase 24 60 [ubiquinone] 1 beta subcomplex subunit 4 26 46 25 38 NADH dehydrogenase 44 31 [ubiquinone] 1 alpha subcomplex subunit 5 44 33 44 32 NADH dehydrogenase 69 32 [ubiquinone] 1 alpha subcomplex subunit 6 66 33 67 33 NADH dehydrogenase 56 80 [ubiquinone] iron-sulfur protein 7 58 74 53 78 NADH dehydrogenase 52 66 [ubiquinone] 1 alpha subcomplex subunit 8 52 61 52 57 NADH dehydrogenase 90 44 [ubiquinone] 1 alpha subcomplex subunit 9 88 34 82 37 NADH dehydrogenase 30 47 [ubiquinone] 1 alpha subcomplex subunit 11 30 46 27 41 NADH dehydrogenase 98 44 [ubiquinone] 1 alpha subcomplex subunit 12 98 42 99 45 Electron transport chain complex II Succinate dehydrogenase subunit A 99 69 98 68 99 66 Succinate dehydrogenase subunit B 43 85 43 81 42 82 Electron transport chain complex III Cytochrome b5 85 45 85 44 88 41 Cytochrome b-cl complex subunit Rieske 50 75 58 67 58 63 Cytochrome b-cl complex subunit 1 79 38 79 38 74 37 Cytochrome b-cl complex subunit 2 61 41 61 43 56 42 Cytochrome b-cl complex subunit 7 38 61 38 61 38 59 Cytochrome cl 85 68 82 50 82 49 Electron transport chain complex IV Cytochrome c 26 90 26 82 26 86 COX 7 subunit A 38 36 21 48 21 49 COX 11 48 80 48 78 48 76 COX 15 46 70 46 68 46 62 COX 16 31 65 32 57 30 59 Electron transport chain complex V ATP synthase subunit D 32 83 32 83 32 80 ATP synthase subunit E 46 57 53 53 43 55 ATP synthase subunit F 48 72 48 73 48 71 ATP synthase subunit G 11 82 10 62 11 57 ATP synthase subunit O 50 60 50 58 49 56 Krebs cycle Citrate synthase 42 79 44 74 43 72 Aconitase 70 79 69 79 69 79 Succinyl-CoA synthetase 57 78 subunit alpha 51 84 58 78 ADP-forming Succinyl-CoA 69 68 ligase subunit beta 69 67 69 67 Mitochondrial NADP(+)-dependent 52 78 Isocitrate dehydrogenase 55 70 54 72 Mitochondrial NAD(+)-Isocitrate 67 73 dehydrogenase subunit alpha 67 75 64 75 Mitochondrial NAD(+)-Isocitrate 12 66 dehydrogenase subunit beta 12 65 12 67 Mitochondrial NAD(+)-Isocitrate 53 68 dehydrogenase subunit gamma 57 64 53 67 Malate dehydrogenase 72 77 72 76 71 77 Fumarase 65 83 65 79 68 77 2-Oxoglutarate dehydrogenase 85 76 85 76 89 72 Dihydrolipoyllysine-residue 29 79 succinyltransferase 25 78 30 78 Pyruvate dehydrogenase complex Pyruvate dehydrogenase El 79 50 component subunit alpha 78 56 72 55 Pyruvate dehydrogenase El 31 82 component subunit beta 32 75 32 79 Dihydrolipoyllysine-residue 99 59 acetyltransferase 99 59 99 61 Dihydrolipoyl dehydrogenase 60 75 60 75 60 74 Mitochondrial carriers, permeability transition, and calcium transport proteins Voltage-dependent anion channel 38 94 38 63 38 61 Adenine nucleotide 90 59 translocator isoform 3 92 58 91 59 Cyclophilin D 28 56 2S 53 2S 55 Phosphate carrier protein 61 75 61 74 61 73 All sequences are encoded in the nuclear genome. The second column contains the transcript or expressed sequence tags Genebank access number previously reported for the protein analyzed in other species of peneids.