**1. Introduction**

#### **1.1. Relevance of carbohydrate metabolism studies in fish**

The zebrafish *Danio rerio* has long been used as an animal model for developmental studies due to a number of desirable characteristics for lab study including its short generation time, large number of offspring, transparent embryos, and *ex utero* development of the embryo. This setting proved to be particularly useful for research on vertebrate development [1] and modeling of human disease [2], namely the hematopoietic [3, 4] and the cardiac systems [5, 6]. However, the use of fish for scientific studies has evolved substantially in the last 30 years. The direction of metabolic fish studies changed drastically with the advent of modern aquaculture and a high demand for models to study nutrition, physiology and metabolism. Capture of fish products in the end of the 1990's represented almost 75% of total production but since 2001 that has leveled at around 90 million tons. Meanwhile, aquaculture produc‐ tion has been increasing at an average annual growth rate of 6.2% from 38.9 million tons in 2003 to 52.5 million tons in 2008 [7]. Aquaculture in 2009 already accounted for 38% of the 145 million tones of total fish products, 81% of which were for human consumption [8]. It is estimated that by 2030, half of the production for human consumption will be derived from aquaculture while the harvest of wild fish will not show any significant growth [9]. Farming of aquatic species has existed for many thousands of years: in ancient Asia, carp were left to grow in ponds and rice paddies and later harvested. Similar practices were thought to take place in ancient Egypt with tilapia and in southern Europe in a polyculture regime, includ‐ ing mullet *Mugil* spp, sole *Solea* spp, seabass *Dicentrarchus labrax* and gilthead seabream *Spa‐ rus aurata* [10]. These artisanal methods, with little or no active manipulation of the animals

© 2013 Viegas et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 Viegas et al.; licensee InTech. This is a paper distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

or diets, are still practiced in some parts of the globe. However, as it has evolved into a high‐ ly competitive and commercialized business, the technique of aquaculture has also signifi‐ cantly evolved both in targeted species and in farming methodologies. Aquaculture is highly dependent on capture fisheries to provide fishmeal and fish oil required to produce feeds [11, 12]. Thus, the development of well-suited and cost-effective substitute feeds based on carbohydrates has become a matter of extreme importance to the sustainability and prof‐ itability of the sector. Along with it, comes the need to improve our understanding of fish carbohydrate nutrition, its physiology, biochemistry and metabolism.

glucose anabolism. As for mammals, many tissues such as skeletal muscle can oxidize alter‐ native substrates to glucose for energy such as triglycerides and can therefore function with variable levels of blood glucose [20] but the brain is highly dependent on glucose as an ener‐ gy substrate therefore a threshold level of plasma glucose is required for maintaining its function. Not surprisingly, the brain is regarded as key component of endogenous glucose sensing mechanisms that have recently been described in fish models [21]. The plasma glu‐ cose concentration also has considerable influence on the selection of myocardial substrates for oxidation in resting and active fish [22] and this adaptation is representative of the changes that occur in whole-body glucose utilization in response to changes in glucose availability [23]. During hypoxia, tissues such as the heart and brain upregulate anaerobic metabolic pathways and become more reliant on glycolysis for ATP production [24]. How‐ ever under normoxic conditions, glucose uptake by muscle is surpassed by that of amino acids and lipids for growth and energy [25]; this also happens during the recovery of muscle glycogen following exhaustive exercise. Thus in general, the direct contribution of blood glucose to muscle energy metabolism seems to be minor [20, 26, 27, 28, 29]. The generally low reliance of fish muscle on glucose as an energy substrate is compatible with the limited

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In mammals, glucose enters the hepatocyte through the GLUT-2 transporter, which is pri‐ marily expressed in the liver but is also present in kidney, intestine and β-cells of pancreatic islets [19]. Due to its low affinity and high capacity, GLUT-2 transports hexoses in a large range of physiological concentrations necessary for glucose homeostasis, displaying bidirec‐ tional fluxes in and out the cells, namely in hepatocytes [30]. Recently, the presence of GLUT-2 transporter was confirmed at both biochemical and molecular levels in species like rainbow trout *Oncorhynchus mykiss* [31, 32] and its expression in seabass *D. labrax* hepato‐

Blood glucose concentration is the net result of the difference between rates of glucose ap‐ pearance and disposal. Following uptake into the hepatocyte, the ATP-dependent conver‐ sion of glucose to glucose 6-phosphate (G6P) is catalyzed by hexokinase IV or glucokinase (GK). GK is the major hexokinase expressed in liver and due to its relatively high Km, its role in glucose homeostasis, is to increase the hepatic capacity for glucose metabolism during hy‐ perglycemia. Uniquely, GK is not inhibited by its G6P product so it is able to maintain high rates of glucose phosphorylation while the activities of other hexokinases become limited by increased G6P levels. Moreover, GK activity is hormonally controlled via the glucokinase regulatory protein (GKRP), such that its activity is increased by the high insulin levels of the fed state, but rapidly suppressed during the onset of fasting when both portal vein glucose and insulin levels are waning. As a result, when the liver becomes a net producer of glucose

The G6P product can be utilized by the pentose phosphate pathway, be recruited for glyco‐ gen synthesis (glycogenesis) or be metabolized by the glycolytic pathway to pyruvate. Pyru‐ vate in turn may be oxidized to acetyl coenzyme A (acetyl-CoA) or carboxylated to

during the fasting state, GK is prevented from converting this back to G6P.

availability of carbohydrate in fish diets [18].

**1.3. Hepatic glucose storage and disposal**

cytes seems to be regulated by hypoxia.

#### **1.2. Carbohydrate metabolism in carnivorous fish**

The central organization of metabolic pathways is highly conserved amongst vertebrates. While the metabolic machinery of fish is much the same as that of mammals, the main dif‐ ferences lie in the nutritional and endocrine control of the pathways via various feedback mechanisms. As a water-living organism, fish have specific adaptations in relation to their terrestrial vertebrate counter-parts. As consequence of living in water, their constant ther‐ mal equilibrium with the environment is a principal determinant of overall metabolic rates while excess dietary nitrogen can be cleared as ammonia, a highly toxic but rapidly diffusi‐ ble molecule that is efficiently transferred from blood to the outside water via the gills [13]. Carbohydrates (CHO) are a basic nutritional source of energy and carbon, but carnivorous fish have a limited capability to digest and metabolize them since they are adapted to a diet high in protein [14]. CHO influence growth, feed utilization and deposition of nutrients ac‐ cording to species, quantity, origin and treatment of dietary CHO used [15, 16, 17]. Fish are considered to be glucose intolerant on the basis of sluggish clearance of a glucose load and their metabolism of glucose has often been compared to that of mammals with insulin-de‐ pendent diabetes *mellitus* (IDDM) [18]. However, unlike IDDM, the causes of this intoler‐ ance cannot be attributed to a simple deficiency of insulin but is instead a reflection of different enzymatic and hormonal control regimes for glucose regulation in fish compared to mammals. The liver utilizes, produces and stores CHO and is an important component of systemic glycemic control in vertebrates. The rate of nutrient absorption coupled with con‐ tinuous monitoring of blood glucose levels by various organs including the brain and pan‐ creas mediates a combination of nutritional, endocrine and nerve-mediated regulation of hepatic glucose metabolism. This results in net hepatic glucose uptake when CHO is abun‐ dant, such as after feeding and net hepatic glucose output when dietary CHO is unavailable, such as during starvation. Glucose is the major energy source for the central nervous system and the only energy source for erythrocytes and the functions of both are dependent on a threshold concentration of blood glucose. Blood glucose levels are maintained through a balance of several factors, including the rate of consumption and intestinal absorption of di‐ etary CHO, the rate of utilization of glucose by peripheral tissues and the loss of glucose through the kidney and finally the rate of removal or release of glucose by the liver [19].

Figure 1 depicts the pathways by which the liver disposes glucose into the bloodstream dur‐ ing fasting and storage and utilization of imported glucose during feeding. The main en‐ zymes involved in these pathways are also represented as well as possible final fates of

glucose anabolism. As for mammals, many tissues such as skeletal muscle can oxidize alter‐ native substrates to glucose for energy such as triglycerides and can therefore function with variable levels of blood glucose [20] but the brain is highly dependent on glucose as an ener‐ gy substrate therefore a threshold level of plasma glucose is required for maintaining its function. Not surprisingly, the brain is regarded as key component of endogenous glucose sensing mechanisms that have recently been described in fish models [21]. The plasma glu‐ cose concentration also has considerable influence on the selection of myocardial substrates for oxidation in resting and active fish [22] and this adaptation is representative of the changes that occur in whole-body glucose utilization in response to changes in glucose availability [23]. During hypoxia, tissues such as the heart and brain upregulate anaerobic metabolic pathways and become more reliant on glycolysis for ATP production [24]. How‐ ever under normoxic conditions, glucose uptake by muscle is surpassed by that of amino acids and lipids for growth and energy [25]; this also happens during the recovery of muscle glycogen following exhaustive exercise. Thus in general, the direct contribution of blood glucose to muscle energy metabolism seems to be minor [20, 26, 27, 28, 29]. The generally low reliance of fish muscle on glucose as an energy substrate is compatible with the limited availability of carbohydrate in fish diets [18].

#### **1.3. Hepatic glucose storage and disposal**

or diets, are still practiced in some parts of the globe. However, as it has evolved into a high‐ ly competitive and commercialized business, the technique of aquaculture has also signifi‐ cantly evolved both in targeted species and in farming methodologies. Aquaculture is highly dependent on capture fisheries to provide fishmeal and fish oil required to produce feeds [11, 12]. Thus, the development of well-suited and cost-effective substitute feeds based on carbohydrates has become a matter of extreme importance to the sustainability and prof‐ itability of the sector. Along with it, comes the need to improve our understanding of fish

The central organization of metabolic pathways is highly conserved amongst vertebrates. While the metabolic machinery of fish is much the same as that of mammals, the main dif‐ ferences lie in the nutritional and endocrine control of the pathways via various feedback mechanisms. As a water-living organism, fish have specific adaptations in relation to their terrestrial vertebrate counter-parts. As consequence of living in water, their constant ther‐ mal equilibrium with the environment is a principal determinant of overall metabolic rates while excess dietary nitrogen can be cleared as ammonia, a highly toxic but rapidly diffusi‐ ble molecule that is efficiently transferred from blood to the outside water via the gills [13]. Carbohydrates (CHO) are a basic nutritional source of energy and carbon, but carnivorous fish have a limited capability to digest and metabolize them since they are adapted to a diet high in protein [14]. CHO influence growth, feed utilization and deposition of nutrients ac‐ cording to species, quantity, origin and treatment of dietary CHO used [15, 16, 17]. Fish are considered to be glucose intolerant on the basis of sluggish clearance of a glucose load and their metabolism of glucose has often been compared to that of mammals with insulin-de‐ pendent diabetes *mellitus* (IDDM) [18]. However, unlike IDDM, the causes of this intoler‐ ance cannot be attributed to a simple deficiency of insulin but is instead a reflection of different enzymatic and hormonal control regimes for glucose regulation in fish compared to mammals. The liver utilizes, produces and stores CHO and is an important component of systemic glycemic control in vertebrates. The rate of nutrient absorption coupled with con‐ tinuous monitoring of blood glucose levels by various organs including the brain and pan‐ creas mediates a combination of nutritional, endocrine and nerve-mediated regulation of hepatic glucose metabolism. This results in net hepatic glucose uptake when CHO is abun‐ dant, such as after feeding and net hepatic glucose output when dietary CHO is unavailable, such as during starvation. Glucose is the major energy source for the central nervous system and the only energy source for erythrocytes and the functions of both are dependent on a threshold concentration of blood glucose. Blood glucose levels are maintained through a balance of several factors, including the rate of consumption and intestinal absorption of di‐ etary CHO, the rate of utilization of glucose by peripheral tissues and the loss of glucose through the kidney and finally the rate of removal or release of glucose by the liver [19].

Figure 1 depicts the pathways by which the liver disposes glucose into the bloodstream dur‐ ing fasting and storage and utilization of imported glucose during feeding. The main en‐ zymes involved in these pathways are also represented as well as possible final fates of

carbohydrate nutrition, its physiology, biochemistry and metabolism.

**1.2. Carbohydrate metabolism in carnivorous fish**

248 New Advances and Contributions to Fish Biology

In mammals, glucose enters the hepatocyte through the GLUT-2 transporter, which is pri‐ marily expressed in the liver but is also present in kidney, intestine and β-cells of pancreatic islets [19]. Due to its low affinity and high capacity, GLUT-2 transports hexoses in a large range of physiological concentrations necessary for glucose homeostasis, displaying bidirec‐ tional fluxes in and out the cells, namely in hepatocytes [30]. Recently, the presence of GLUT-2 transporter was confirmed at both biochemical and molecular levels in species like rainbow trout *Oncorhynchus mykiss* [31, 32] and its expression in seabass *D. labrax* hepato‐ cytes seems to be regulated by hypoxia.

Blood glucose concentration is the net result of the difference between rates of glucose ap‐ pearance and disposal. Following uptake into the hepatocyte, the ATP-dependent conver‐ sion of glucose to glucose 6-phosphate (G6P) is catalyzed by hexokinase IV or glucokinase (GK). GK is the major hexokinase expressed in liver and due to its relatively high Km, its role in glucose homeostasis, is to increase the hepatic capacity for glucose metabolism during hy‐ perglycemia. Uniquely, GK is not inhibited by its G6P product so it is able to maintain high rates of glucose phosphorylation while the activities of other hexokinases become limited by increased G6P levels. Moreover, GK activity is hormonally controlled via the glucokinase regulatory protein (GKRP), such that its activity is increased by the high insulin levels of the fed state, but rapidly suppressed during the onset of fasting when both portal vein glucose and insulin levels are waning. As a result, when the liver becomes a net producer of glucose during the fasting state, GK is prevented from converting this back to G6P.

The G6P product can be utilized by the pentose phosphate pathway, be recruited for glyco‐ gen synthesis (glycogenesis) or be metabolized by the glycolytic pathway to pyruvate. Pyru‐ vate in turn may be oxidized to acetyl coenzyme A (acetyl-CoA) or carboxylated to oxaloacetate or malate – a process known as anaplerosis. Acetyl-CoA may be oxidized to CO2 by the tricarboxylic acid cycle (TCA cycle) or utilized for lipid production (lipogenesis) while anaplerotic products can be utilized for gluconeogenesis. G6P can also be hydrolyzed back to glucose, a reaction catalyzed by glucose 6-phosphatase (G6Pase). In fish, it has been shown that G6Pase is poorly regulated by dietary CHO in comparison to GK [33]. When G6Pase and GK are both active at the same time, glucose and G6P are interconverted in a "futile cycle" that results in the consumption of ATP but no net conversion of glucose to products. This activity has important implications for interpreting the metabolism of certain glucose tracers such as [2-2 H]- or [2-3 H]glucose since it results in disappearance of the label without any net glucose consumption.

**1.4. Glycogen synthesis and hydrolysis**

The liver is the main storage site for glycogen, but tissues such as gills, kidney and brain also sustain relatively high rates of glycogen synthesis from glucose. Net glycogen synthesis results from the glycogen synthase (GSase) and glycogen phosphorylase (GPase) activities

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GSase exists in a phosphorylated inactive form in the cytosol and it is activated by binding to G6P and this causes a conformational change that makes it a better substrate for protein phosphatases, which then convert the enzyme to the active dephosphorylated isoform. In glycogenesis, G6P is first converted to glucose 1-phosphate (G1P) by phosphoglucomutase and subsequently to uridine diphosphate glucose (UDPG) by UDPG pyrophosphorylase. Glycogen synthase then adds glucose residues from the UDPG donor to the growing glyco‐ gen molecule via α-1,4 glycosidic linkages. Hepatic glycogen can be synthesized by the in‐ corporation of glucose into glycogen via a sequence of reactions that was initially characterized in muscle. In this so-called "direct pathway", the glucosyl moiety is transfer‐ red into glycogen as an intact entity. It was subsequently discovered that in the liver, glyco‐ gen can also be synthesized independently from glucose via gluconeogenic precursors [19]. These contributions are collectively referred to as the "indirect pathway. Since the indirect pathway bypasses GK, and uses precursors that are independent of blood glucose, its contri‐ bution to glycogen synthesis may respond to nutritional and hormonal states in a different way to that of the direct pathway. Quantifying the direct and indirect pathway flux compo‐ nents of hepatic glycogen synthesis can therefore further inform adaptations of hepatic car‐

Under fasting conditions, the liver becomes a net producer of glucose and the hydrolysis of glycogen is an important source of glucose production at least in the early stages of fasting. To avoid futile cycling between G6P and glycogen, the activities of GSase and GPase are regulated in a reciprocal manner such that only one or the other is active at a given time. Glycogen may be also hydrolyzed to glucose by the action of glucosidases enzymes that hy‐ drolyze the non-reducing end of polysaccharides directly to glucose. There is evidence of glucosidase activity alongside that of GPase in living fish [35, 36, 37], but the extent of its contribution to glucose synthesis from glycogen and modulation of activity by feeding and

Most fish species need to periodically cope with starvation as a result of seasonal food limi‐ tation or in some cases as natural consequence of their life cycle [38] and the liver plays an important role in the endogenous control of fuel storage and mobilization [39, 40, 41]. Glyco‐ gen represents an important source of glucose to be released in the fasted state or when re‐ sponding to acute stressors. The transition from fasting to feeding states results in a comprehensive realignment of hepatic carbohydrate metabolic fluxes from minimal mainte‐ nance of glycemia and peripheral glucose demands by endogenous glucose production dur‐ ing the fasting phase to a high nutrient inflow and replenishment of liver glycogen stores during refeeding [39, 40, 42, 43]. The general consensus is that the liver of fish plays a role in glycemic control through the regulation of hepatic glucose storage and mobilization [33] but

that are reciprocally regulated both allosterically and by phosphorylation [34].

bohydrate metabolism to different diets and hormonal states.

its actions are sluggish in comparison to mammalian liver.

fasting are little known.

**Figure 1.** Metabolic model representing main pathways in the liver involving catabolism and anabolism of glucose. Gluconeogenic precursors are represented by pyruvate (and gluconeogenic amino acids, metabolized via the anapler‐ otic pathways of TCA cycle) as well as glycerol from lipolysis. Some metabolic intermediates were omitted for clarity. Abbreviations are as follows: G6P - glucose 6-phosphate; G1P - glucose 1-phosphate; UDPG - uridine diphosphoglu‐ cose; F6P - fructose 6-phosphate; F16P2 - fructose-1,6-bisphosphate; G3P - glyceraldehyde 3-phosphate; DHAP - dihy‐ droxyacetone phosphate; PEP - phosphoenolpyruvate; TG's - triglycerides; OA - oxaloacetate; AA's - amino acids; GK glucokinase; G6Pase - glucose-6-phosphatase; FBPase - fructose bisphosphatase; PFK-1 - 6-phosphofructo-1-kinase; PEPCK - phosphoenolpyruvate carboxykinase; PK - pyruvate kinase; GPase - glycogen phosphatase; Glycosid - glycosi‐ dic enzymes; TCA cycle - tricarboxylic acid cycle.

#### **1.4. Glycogen synthesis and hydrolysis**

oxaloacetate or malate – a process known as anaplerosis. Acetyl-CoA may be oxidized to CO2 by the tricarboxylic acid cycle (TCA cycle) or utilized for lipid production (lipogenesis) while anaplerotic products can be utilized for gluconeogenesis. G6P can also be hydrolyzed back to glucose, a reaction catalyzed by glucose 6-phosphatase (G6Pase). In fish, it has been shown that G6Pase is poorly regulated by dietary CHO in comparison to GK [33]. When G6Pase and GK are both active at the same time, glucose and G6P are interconverted in a "futile cycle" that results in the consumption of ATP but no net conversion of glucose to products. This activity has important implications for interpreting the metabolism of certain

**Figure 1.** Metabolic model representing main pathways in the liver involving catabolism and anabolism of glucose. Gluconeogenic precursors are represented by pyruvate (and gluconeogenic amino acids, metabolized via the anapler‐ otic pathways of TCA cycle) as well as glycerol from lipolysis. Some metabolic intermediates were omitted for clarity. Abbreviations are as follows: G6P - glucose 6-phosphate; G1P - glucose 1-phosphate; UDPG - uridine diphosphoglu‐ cose; F6P - fructose 6-phosphate; F16P2 - fructose-1,6-bisphosphate; G3P - glyceraldehyde 3-phosphate; DHAP - dihy‐ droxyacetone phosphate; PEP - phosphoenolpyruvate; TG's - triglycerides; OA - oxaloacetate; AA's - amino acids; GK glucokinase; G6Pase - glucose-6-phosphatase; FBPase - fructose bisphosphatase; PFK-1 - 6-phosphofructo-1-kinase; PEPCK - phosphoenolpyruvate carboxykinase; PK - pyruvate kinase; GPase - glycogen phosphatase; Glycosid - glycosi‐

H]glucose since it results in disappearance of the label

glucose tracers such as [2-2

250 New Advances and Contributions to Fish Biology

without any net glucose consumption.

dic enzymes; TCA cycle - tricarboxylic acid cycle.

H]- or [2-3

The liver is the main storage site for glycogen, but tissues such as gills, kidney and brain also sustain relatively high rates of glycogen synthesis from glucose. Net glycogen synthesis results from the glycogen synthase (GSase) and glycogen phosphorylase (GPase) activities that are reciprocally regulated both allosterically and by phosphorylation [34].

GSase exists in a phosphorylated inactive form in the cytosol and it is activated by binding to G6P and this causes a conformational change that makes it a better substrate for protein phosphatases, which then convert the enzyme to the active dephosphorylated isoform. In glycogenesis, G6P is first converted to glucose 1-phosphate (G1P) by phosphoglucomutase and subsequently to uridine diphosphate glucose (UDPG) by UDPG pyrophosphorylase. Glycogen synthase then adds glucose residues from the UDPG donor to the growing glyco‐ gen molecule via α-1,4 glycosidic linkages. Hepatic glycogen can be synthesized by the in‐ corporation of glucose into glycogen via a sequence of reactions that was initially characterized in muscle. In this so-called "direct pathway", the glucosyl moiety is transfer‐ red into glycogen as an intact entity. It was subsequently discovered that in the liver, glyco‐ gen can also be synthesized independently from glucose via gluconeogenic precursors [19]. These contributions are collectively referred to as the "indirect pathway. Since the indirect pathway bypasses GK, and uses precursors that are independent of blood glucose, its contri‐ bution to glycogen synthesis may respond to nutritional and hormonal states in a different way to that of the direct pathway. Quantifying the direct and indirect pathway flux compo‐ nents of hepatic glycogen synthesis can therefore further inform adaptations of hepatic car‐ bohydrate metabolism to different diets and hormonal states.

Under fasting conditions, the liver becomes a net producer of glucose and the hydrolysis of glycogen is an important source of glucose production at least in the early stages of fasting. To avoid futile cycling between G6P and glycogen, the activities of GSase and GPase are regulated in a reciprocal manner such that only one or the other is active at a given time. Glycogen may be also hydrolyzed to glucose by the action of glucosidases enzymes that hy‐ drolyze the non-reducing end of polysaccharides directly to glucose. There is evidence of glucosidase activity alongside that of GPase in living fish [35, 36, 37], but the extent of its contribution to glucose synthesis from glycogen and modulation of activity by feeding and fasting are little known.

Most fish species need to periodically cope with starvation as a result of seasonal food limi‐ tation or in some cases as natural consequence of their life cycle [38] and the liver plays an important role in the endogenous control of fuel storage and mobilization [39, 40, 41]. Glyco‐ gen represents an important source of glucose to be released in the fasted state or when re‐ sponding to acute stressors. The transition from fasting to feeding states results in a comprehensive realignment of hepatic carbohydrate metabolic fluxes from minimal mainte‐ nance of glycemia and peripheral glucose demands by endogenous glucose production dur‐ ing the fasting phase to a high nutrient inflow and replenishment of liver glycogen stores during refeeding [39, 40, 42, 43]. The general consensus is that the liver of fish plays a role in glycemic control through the regulation of hepatic glucose storage and mobilization [33] but its actions are sluggish in comparison to mammalian liver.

#### **1.5. Glycolysis and gluconeogenesis**

Glycolysis involves the metabolism of glucose or glycogen to pyruvate. From glucose,the first step in the glycolytic pathway is the phosphorylation of glucose by GK into G6P, as descri‐ bed. From glycogen, G6P is generated following GPase and phosphoglucomutase activities. G6P is isomerized to F6P via G6P isomerase and this is followed by another phosphorylation of fructose 6-phosphate (F6P) to fructose 1,6-bisphosphate (F16P2) catalyzed by 6-phospho‐ fructo 1-kinase (PFK-1). These reactions commit the hexose carbon skeletons to pyruvate pro‐ duction. Both phosphorylations are highly regulated and along with conversion of phosphoenolpyruvate (PEP) to pyruvate catalyzed by pyruvate kinase (PK) constitute the ir‐ reversible steps of glycolysis. The energy charge of the cell, as well as allosteric and transcrip‐ tional processes, contributes to control fluxes through PFK-1 and PK. The free energy released in this process is coupled to substrate-level ATP and NADH formation. Pyruvate is the final product of glycolysis and can undergo further oxidation to acetyl-CoA via pyruvate dehydro‐ genase (PDH), carboxylation via pyruvate carboxylase (PC) or NADP-malic enzyme or reduc‐ tion to lactate. In the liver, the relative activities of PDH and PC determine whether pyruvate is oxidized or utilized by anaplerotic pathways such as gluconeogensis hence the activities of these enzymes are highly regulated depending on the nutritional state. Gluconeogenesis is the principal metabolic pathway that generates glucose in fish. Its main precursors include lactate (converted into pyruvate by lactate dehydrogenase) and gluconeogenic amino acids (those that are metabolized to pyruvate or C4 and C5 TCA cycle intermediates), with minor contributions from glycerol, derived by lipolysis of triglycerides. While conversion of pyru‐ vate to glucose occurs via the same intermediates as glycolysis, different enzymes are used to overcome the unfavorable free energy change of pyruvate to PEP conversion. The conversion of F16P2 to F6P and G6P to glucose are also mediated by different enzymes to their glycolytic counter-parts (fructose 1,6-bisphosphatase and glucose 6-phosphatase, respectively). Pyru‐ vate is converted to PEP at the expense of two ATP equivalents. First, pyruvate carboxylase generates oxaloacetate (OA) from pyruvate and second, PEP carboxykinase (PEPCK) con‐ verts OA to PEP. The free energy change of pyruvate conversion to glucose is positive, hence pyruvate gluconeogenesis requires energy in the form of ATP and reducing equivalents. This is not the case for all gluconeogenic precursors, notably glutamate and glutamine. These are metabolized to α-ketoglutarate, which during its oxidation to OA via the TCA cycle, gener‐ ates ATP and reducing equivalents in excess of those consumed by PEPCK and glyceralde‐ hyde 3-phosphate dehydrogenase. It is important to note that these amino acids are among the most abundant in dietary protein, therefore from a thermodynamic viewpoint, they facili‐ tate gluconeogenesis from protein.

Hormones in vertebrate organisms are a diverse but well conserved group of signaling mol‐ ecules that regulate and modulate metabolic fluxes. The role of hormones in the regulation of CHO metabolism of fish are far from being completely known due to the variety of re‐ sponses observed - many of which are dependent on whether studies are performed on in‐ tact fish, isolated organs or primary cell cultures. The mammalian paradigm, to which the hormonal regulation in fish is usually compared, has shown to be poorly representative of fish metabolism in a number of cases [62]. Glucose is weakly effective at stimulating insulin secretion in fish compared to mammalian species, although, as expected, an improved re‐ sponse is observed in CHO-tolerant species [63]. On the other hand, amino acids are potent stimulators of insulin secretion even though the intensity of response varies greatly between salmonids and other fish like carps and seabream. This bears some resemblance to mam‐

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In mammals, the actions of insulin are directly opposed by glucagon and adrenal hormones such as adrenaline and these counterregulatory actions are an important component of glu‐ cose homeostasis. However, it is unclear in fish to what extent the regulatory and counterre‐ gulatory processes are coupled [62]. For example, in response to stress or hypoglycemia, the adrenaline-induced hyperglycemic response of rainbow trout is caused by the stimulation of hepatic glucose production that happens in a dose-dependent fashion [64]. However, the ac‐ companying suppression of glucose clearance described in mammals is not observed in fish. Cortisol is a major regulator of intermediary metabolism and promotes hepatic glucose pro‐ duction and hyperglycemia, primarily as a result of increased hepatic gluconeogenesis fuel‐

A metabolic tracer is, by definition, a substance used to follow the biological transformation of an endogenous substrate (tracee). The tracer must have a unique property that allows its detection but at the same time be chemically identical to the tracee. In most cases, tracers consist of synthetic substrate molecules where one or more atoms in the molecule are substi‐ tuted for an atom of the same chemical element, but of a different isotope. Isotopic tracers may be identified by radioactivity or differences in mass and/or nuclear spin from the tracee. In order that the tracee pool of interest be not altered by administration of the tracer (thereby potentially perturbing the metabolic activity under observation), its chemical con‐ centration should be insignificant in comparison to that of the tracee. At the same time, this amount of material must possess a high abundance of label relative to the tracee since meta‐ bolic flux parameters are typically derived by administering a high density of label and monitoring its dilution by unlabeled tracee molecules. With radioisotopes, the label concen‐ tration is measured in terms of specific activity, defined as the number of decays per minute measured per mol of glucose. With stable isotopes, tracer concentration is defined in terms of percent excess enrichment per mol of glucose. Since most stable isotopes are typically

mals, where arginine is also a powerful insulin secretagogue.

led in part by amino acid products of peripheral proteolysis [65].

**2. Tracer studies of carbohydrate metabolism in fish**

**2.1. Overview**

#### **1.6. Enzymatic and hormonal regulation**

Measurement of enzymatic activities involved directly or indirectly in CHO metabolism has been performed in various species and tissues and has proved to be of crucial importance in evaluating adaptation to changes in temperature [44, 45], salinity and osmoregulation [46, 47], feeding status [39, 41, 48, 49], rearing densities [50], diets [51, 52, 53, 54, 55, 56, 57] and to study the effects of hormones [58, 59, 60, 61].

Hormones in vertebrate organisms are a diverse but well conserved group of signaling mol‐ ecules that regulate and modulate metabolic fluxes. The role of hormones in the regulation of CHO metabolism of fish are far from being completely known due to the variety of re‐ sponses observed - many of which are dependent on whether studies are performed on in‐ tact fish, isolated organs or primary cell cultures. The mammalian paradigm, to which the hormonal regulation in fish is usually compared, has shown to be poorly representative of fish metabolism in a number of cases [62]. Glucose is weakly effective at stimulating insulin secretion in fish compared to mammalian species, although, as expected, an improved re‐ sponse is observed in CHO-tolerant species [63]. On the other hand, amino acids are potent stimulators of insulin secretion even though the intensity of response varies greatly between salmonids and other fish like carps and seabream. This bears some resemblance to mam‐ mals, where arginine is also a powerful insulin secretagogue.

In mammals, the actions of insulin are directly opposed by glucagon and adrenal hormones such as adrenaline and these counterregulatory actions are an important component of glu‐ cose homeostasis. However, it is unclear in fish to what extent the regulatory and counterre‐ gulatory processes are coupled [62]. For example, in response to stress or hypoglycemia, the adrenaline-induced hyperglycemic response of rainbow trout is caused by the stimulation of hepatic glucose production that happens in a dose-dependent fashion [64]. However, the ac‐ companying suppression of glucose clearance described in mammals is not observed in fish. Cortisol is a major regulator of intermediary metabolism and promotes hepatic glucose pro‐ duction and hyperglycemia, primarily as a result of increased hepatic gluconeogenesis fuel‐ led in part by amino acid products of peripheral proteolysis [65].

## **2. Tracer studies of carbohydrate metabolism in fish**

#### **2.1. Overview**

**1.5. Glycolysis and gluconeogenesis**

252 New Advances and Contributions to Fish Biology

tate gluconeogenesis from protein.

**1.6. Enzymatic and hormonal regulation**

study the effects of hormones [58, 59, 60, 61].

Glycolysis involves the metabolism of glucose or glycogen to pyruvate. From glucose,the first step in the glycolytic pathway is the phosphorylation of glucose by GK into G6P, as descri‐ bed. From glycogen, G6P is generated following GPase and phosphoglucomutase activities. G6P is isomerized to F6P via G6P isomerase and this is followed by another phosphorylation of fructose 6-phosphate (F6P) to fructose 1,6-bisphosphate (F16P2) catalyzed by 6-phospho‐ fructo 1-kinase (PFK-1). These reactions commit the hexose carbon skeletons to pyruvate pro‐ duction. Both phosphorylations are highly regulated and along with conversion of phosphoenolpyruvate (PEP) to pyruvate catalyzed by pyruvate kinase (PK) constitute the ir‐ reversible steps of glycolysis. The energy charge of the cell, as well as allosteric and transcrip‐ tional processes, contributes to control fluxes through PFK-1 and PK. The free energy released in this process is coupled to substrate-level ATP and NADH formation. Pyruvate is the final product of glycolysis and can undergo further oxidation to acetyl-CoA via pyruvate dehydro‐ genase (PDH), carboxylation via pyruvate carboxylase (PC) or NADP-malic enzyme or reduc‐ tion to lactate. In the liver, the relative activities of PDH and PC determine whether pyruvate is oxidized or utilized by anaplerotic pathways such as gluconeogensis hence the activities of these enzymes are highly regulated depending on the nutritional state. Gluconeogenesis is the principal metabolic pathway that generates glucose in fish. Its main precursors include lactate (converted into pyruvate by lactate dehydrogenase) and gluconeogenic amino acids (those that are metabolized to pyruvate or C4 and C5 TCA cycle intermediates), with minor contributions from glycerol, derived by lipolysis of triglycerides. While conversion of pyru‐ vate to glucose occurs via the same intermediates as glycolysis, different enzymes are used to overcome the unfavorable free energy change of pyruvate to PEP conversion. The conversion of F16P2 to F6P and G6P to glucose are also mediated by different enzymes to their glycolytic counter-parts (fructose 1,6-bisphosphatase and glucose 6-phosphatase, respectively). Pyru‐ vate is converted to PEP at the expense of two ATP equivalents. First, pyruvate carboxylase generates oxaloacetate (OA) from pyruvate and second, PEP carboxykinase (PEPCK) con‐ verts OA to PEP. The free energy change of pyruvate conversion to glucose is positive, hence pyruvate gluconeogenesis requires energy in the form of ATP and reducing equivalents. This is not the case for all gluconeogenic precursors, notably glutamate and glutamine. These are metabolized to α-ketoglutarate, which during its oxidation to OA via the TCA cycle, gener‐ ates ATP and reducing equivalents in excess of those consumed by PEPCK and glyceralde‐ hyde 3-phosphate dehydrogenase. It is important to note that these amino acids are among the most abundant in dietary protein, therefore from a thermodynamic viewpoint, they facili‐

Measurement of enzymatic activities involved directly or indirectly in CHO metabolism has been performed in various species and tissues and has proved to be of crucial importance in evaluating adaptation to changes in temperature [44, 45], salinity and osmoregulation [46, 47], feeding status [39, 41, 48, 49], rearing densities [50], diets [51, 52, 53, 54, 55, 56, 57] and to

A metabolic tracer is, by definition, a substance used to follow the biological transformation of an endogenous substrate (tracee). The tracer must have a unique property that allows its detection but at the same time be chemically identical to the tracee. In most cases, tracers consist of synthetic substrate molecules where one or more atoms in the molecule are substi‐ tuted for an atom of the same chemical element, but of a different isotope. Isotopic tracers may be identified by radioactivity or differences in mass and/or nuclear spin from the tracee. In order that the tracee pool of interest be not altered by administration of the tracer (thereby potentially perturbing the metabolic activity under observation), its chemical con‐ centration should be insignificant in comparison to that of the tracee. At the same time, this amount of material must possess a high abundance of label relative to the tracee since meta‐ bolic flux parameters are typically derived by administering a high density of label and monitoring its dilution by unlabeled tracee molecules. With radioisotopes, the label concen‐ tration is measured in terms of specific activity, defined as the number of decays per minute measured per mol of glucose. With stable isotopes, tracer concentration is defined in terms of percent excess enrichment per mol of glucose. Since most stable isotopes are typically present at background levels (for example, 1.1% of carbon in nature is 13C), this must be ac‐ counted for when attributing metabolite enrichment to inflow from a tracer.

prove our overall understanding of piscine glycemic control, endogenous and exogenous

Advances and Applications of Tracer Measurements of Carbohydrate Metabolism in Fish

mation characterizing the main pathways of hepatic carbohydrate metabolism. They are easily detected by scintillation counting and their background radioactivity is very low in comparison to metabolite specific activity. Due to the fact that they are weak α- and β-ray emitters, and are therefore easily contained and have relatively high maximum permissible dosages, these tracers are still used in some human and many animal studies. They are more widely used in cell cultures for substrate uptake studies and isotope-dilution measurements where the small scale of the experiments can be accommodated by reasonable radiation con‐ tainment measures. For this same reason, radioactive isotopes have been particularly useful in the study of larval nutrition. A supply of good quality fish fry is essential for a successful production of juveniles however knowledge of their nutritional requirements is often quali‐ tative rather than quantitative [70]. In this context the innovative methodologies of incorpo‐ ration of 14C-labeled amino acids [71] or fatty acids [72] in larvae live prey *Artemia* sp. nauplii has improved our understanding of larval nutrition. However, for conventional nu‐ tritional studies involving larger fish and tanks, containment of radioactive tracers is often not practically possible therefore in these settings, their stable isotope counterparts (13C and

H) are typically deployed. Methods for obtaining glucose rates of appearance (Ra) that were

adapted for stable isotope studies. Moreover, stable isotope tracers can potentially yield much richer metabolic information than their radioactive counterparts due to advances in

Tracer measurements of whole-body carbohydrate metabolism were originally developed in humans and experimental animal models such as rodents, dogs and primates. Translating these methodologies to fish presents some additional challenges, the most important being the following: firstly, most mammals have accessible peripheral veins that can be catheter‐ ized for tracer administration and blood sampling. Catheters may be installed temporarily under anesthesia for the duration of the tracer study, or as indwelling entities for longitudi‐ nal studies of conscious free-moving animals. In fish, the dorsal aorta proximal to the gill arches is the main accessible vessel for tracer studies. It has been successfully cannulated for tracer infusion and blood sampling in undisturbed, non-anaesthetized rainbow trout of 800-1200 grams in weight [77, 78, 79]. However, it is unclear if this method can be easily ap‐ plied to smaller sized fish, or to fish with a different gill-body configuration to rainbow trout, for example flatfish. The second major impediment for fish carbohydrate tracer stud‐ ies is the fact that glucose appearance and turnover rates are much slower in comparison to mammals, necessitating far longer infusion periods for achieving isotopic steady-state con‐

positional and isotopomer analysis of metabolite enrichment from 13C and 2

**2.3. Tracer administration and measurements in fish compared to mammals**

and beyond those which are possible for radiolabeled tracers.

H- and 14C-labeled glucose [23, 73, 74, 75, 76] can be easily

H has provided valuable infor‐

http://dx.doi.org/10.5772/54053

255

H that go above

contributions to glucose Ra need to be better defined than they are at present.

**2.2. Radioactive versus stable isotope carbohydrate tracers**

Historically, the use of radioactive isotopes such as 14C and 3

2

originally developed using 3

Since blood glucose and other carbohydrates can be synthesized endogenously as well as absorbed from the diet, several tracer strategies may be applied to inform carbohydrate me‐ tabolism. The first and most easily understood is to incorporate the tracer into dietary carbo‐ hydrate and monitor its rate of appearance in endogenous carbohydrate pools (i.e. plasma glucose and liver glycogen) as well as metabolic endproducts (i.e. respired CO2). This ap‐ proach provides a direct measure of how a specific dietary carbohydrate source is utilized and disposed into endogenous oxidative and nonoxidative pathways. Oral administration of labeled food [66], either by pellets with 13C-labeled starch and 15N-labeled protein [67] or 13C-labeled rotifers [68], and measurement of isotope appearance in endogenous carbohy‐ drate and protein have also been used to measure substrate utilization in fish. In the case of carbohydrate studies, while the dilution of 13C-enrichment between the feed precursor and sampled product metabolite may qualitatively inform the extent of endogenous glucose pro‐ duction (i.e. high dilution equates to high rates of endogenous glucose production relative to absorption of the 13C-enriched carbohydrate), quantitative data rely on knowing the rate of entry of the 13C-enriched dietary carbohydrate into blood glucose. Since this is a function of absorption and cannot be directly measured, this approach provides only a limited in‐ sight into endogenous glucose production. In the second type of measurement, a glucose tracer is directly administered into the blood at a known rate or quantity, and by measuring its dilution by unlabeled endogenous glucose, the rate of appearance (Ra) of blood glucose may be quantitatively defined. Since both absorbed and endogenously-produced glucose contribute to the dilution of the labeled glucose, this measurement does not resolve the con‐ tributions of dietary and endogenous sources. Under fasting conditions where there is no in‐ put of dietary glucose, glucose Ra equates to endogenous glucose production. In mammals and humans, dietary and endogenous components of glucose Ra may be resolved by com‐ bining a feed tracer with a second glucose tracer administered intravenously [69]. The detec‐ tion method must have the capability of resolving the labeling contribution of blood glucose from each tracer. For example, radioactivity from infused [3-3 H]glucose and ingested [U-14C]glucose are resolved by scintillation counters on the basis of their different emission energies. Finally, as previously discussed, endogenous glucose can be derived from multiple sources including gluconeogenesis from a diverse range of precursors as well as glycogen hydrolysis. Neither the labeled dietary carbohydrate nor glucose isotope dilution measure‐ ments inform the sources of endogenous glucose production. This represents a significant limitation for understanding carbohydrate metabolism of carnivorous fish since their diet is low in carbohydrate therefore the plasma glucose rate of appearance (Ra) is likely to domi‐ nated by synthesis from endogenous precursors. Moreover in aquaculture, weaning of car‐ nivorous fish from pure fishmeal to diets supplemented with carbohydrates is an important objective for improved sustainability and reduced environmental impact [18]. Hence, there is continuing interest in understanding to what extent their metabolic phenotype can adapt to increased dietary carbohydrate availability and if gluconeogenic utilization of amino acids is spared under these conditions [16]. To effectively address such questions and to im‐ prove our overall understanding of piscine glycemic control, endogenous and exogenous contributions to glucose Ra need to be better defined than they are at present.

#### **2.2. Radioactive versus stable isotope carbohydrate tracers**

present at background levels (for example, 1.1% of carbon in nature is 13C), this must be ac‐

Since blood glucose and other carbohydrates can be synthesized endogenously as well as absorbed from the diet, several tracer strategies may be applied to inform carbohydrate me‐ tabolism. The first and most easily understood is to incorporate the tracer into dietary carbo‐ hydrate and monitor its rate of appearance in endogenous carbohydrate pools (i.e. plasma glucose and liver glycogen) as well as metabolic endproducts (i.e. respired CO2). This ap‐ proach provides a direct measure of how a specific dietary carbohydrate source is utilized and disposed into endogenous oxidative and nonoxidative pathways. Oral administration of labeled food [66], either by pellets with 13C-labeled starch and 15N-labeled protein [67] or 13C-labeled rotifers [68], and measurement of isotope appearance in endogenous carbohy‐ drate and protein have also been used to measure substrate utilization in fish. In the case of carbohydrate studies, while the dilution of 13C-enrichment between the feed precursor and sampled product metabolite may qualitatively inform the extent of endogenous glucose pro‐ duction (i.e. high dilution equates to high rates of endogenous glucose production relative to absorption of the 13C-enriched carbohydrate), quantitative data rely on knowing the rate of entry of the 13C-enriched dietary carbohydrate into blood glucose. Since this is a function of absorption and cannot be directly measured, this approach provides only a limited in‐ sight into endogenous glucose production. In the second type of measurement, a glucose tracer is directly administered into the blood at a known rate or quantity, and by measuring its dilution by unlabeled endogenous glucose, the rate of appearance (Ra) of blood glucose may be quantitatively defined. Since both absorbed and endogenously-produced glucose contribute to the dilution of the labeled glucose, this measurement does not resolve the con‐ tributions of dietary and endogenous sources. Under fasting conditions where there is no in‐ put of dietary glucose, glucose Ra equates to endogenous glucose production. In mammals and humans, dietary and endogenous components of glucose Ra may be resolved by com‐ bining a feed tracer with a second glucose tracer administered intravenously [69]. The detec‐ tion method must have the capability of resolving the labeling contribution of blood glucose

counted for when attributing metabolite enrichment to inflow from a tracer.

254 New Advances and Contributions to Fish Biology

from each tracer. For example, radioactivity from infused [3-3

[U-14C]glucose are resolved by scintillation counters on the basis of their different emission energies. Finally, as previously discussed, endogenous glucose can be derived from multiple sources including gluconeogenesis from a diverse range of precursors as well as glycogen hydrolysis. Neither the labeled dietary carbohydrate nor glucose isotope dilution measure‐ ments inform the sources of endogenous glucose production. This represents a significant limitation for understanding carbohydrate metabolism of carnivorous fish since their diet is low in carbohydrate therefore the plasma glucose rate of appearance (Ra) is likely to domi‐ nated by synthesis from endogenous precursors. Moreover in aquaculture, weaning of car‐ nivorous fish from pure fishmeal to diets supplemented with carbohydrates is an important objective for improved sustainability and reduced environmental impact [18]. Hence, there is continuing interest in understanding to what extent their metabolic phenotype can adapt to increased dietary carbohydrate availability and if gluconeogenic utilization of amino acids is spared under these conditions [16]. To effectively address such questions and to im‐

H]glucose and ingested

Historically, the use of radioactive isotopes such as 14C and 3 H has provided valuable infor‐ mation characterizing the main pathways of hepatic carbohydrate metabolism. They are easily detected by scintillation counting and their background radioactivity is very low in comparison to metabolite specific activity. Due to the fact that they are weak α- and β-ray emitters, and are therefore easily contained and have relatively high maximum permissible dosages, these tracers are still used in some human and many animal studies. They are more widely used in cell cultures for substrate uptake studies and isotope-dilution measurements where the small scale of the experiments can be accommodated by reasonable radiation con‐ tainment measures. For this same reason, radioactive isotopes have been particularly useful in the study of larval nutrition. A supply of good quality fish fry is essential for a successful production of juveniles however knowledge of their nutritional requirements is often quali‐ tative rather than quantitative [70]. In this context the innovative methodologies of incorpo‐ ration of 14C-labeled amino acids [71] or fatty acids [72] in larvae live prey *Artemia* sp. nauplii has improved our understanding of larval nutrition. However, for conventional nu‐ tritional studies involving larger fish and tanks, containment of radioactive tracers is often not practically possible therefore in these settings, their stable isotope counterparts (13C and 2 H) are typically deployed. Methods for obtaining glucose rates of appearance (Ra) that were originally developed using 3 H- and 14C-labeled glucose [23, 73, 74, 75, 76] can be easily adapted for stable isotope studies. Moreover, stable isotope tracers can potentially yield much richer metabolic information than their radioactive counterparts due to advances in positional and isotopomer analysis of metabolite enrichment from 13C and 2 H that go above and beyond those which are possible for radiolabeled tracers.

#### **2.3. Tracer administration and measurements in fish compared to mammals**

Tracer measurements of whole-body carbohydrate metabolism were originally developed in humans and experimental animal models such as rodents, dogs and primates. Translating these methodologies to fish presents some additional challenges, the most important being the following: firstly, most mammals have accessible peripheral veins that can be catheter‐ ized for tracer administration and blood sampling. Catheters may be installed temporarily under anesthesia for the duration of the tracer study, or as indwelling entities for longitudi‐ nal studies of conscious free-moving animals. In fish, the dorsal aorta proximal to the gill arches is the main accessible vessel for tracer studies. It has been successfully cannulated for tracer infusion and blood sampling in undisturbed, non-anaesthetized rainbow trout of 800-1200 grams in weight [77, 78, 79]. However, it is unclear if this method can be easily ap‐ plied to smaller sized fish, or to fish with a different gill-body configuration to rainbow trout, for example flatfish. The second major impediment for fish carbohydrate tracer stud‐ ies is the fact that glucose appearance and turnover rates are much slower in comparison to mammals, necessitating far longer infusion periods for achieving isotopic steady-state con‐ ditions – a critical requirement for investigating precursor-product relationships of biosyn‐ thetic pathways. This is particularly problematic for quantitative studies of gluconeogenesis based on the delivery of a labeled precursor substrate such as 14C-lactate or alanine, since both the tracee precursor and glucose product pools must be at isotopic equilibrium during the sampling period.

gens, many important precursor metabolites of glucose and glycogen biosynthesis, such as lactate, pyruvate and alanine also become rapidly enriched to the same level as body water for the study duration. Thus, a constant precursor enrichment that can be indefi‐

netics in the catfish *Ictalarus punctatus* [83] and gilthead seabream *S. aurata* [84] as well as blood glucose and hepatic glycogen turnover in seabass *D. labrax* [43, 85]. Because of its enormous potential for informing endogenous carbohydrate metabolism in fish, we

Fish and mammals share common pathways for glucose production and consumption [86]

described and validated for mammals [80, 87, 88] can be applied to fish. Deuterated water

H2O) is a relatively inexpensive non-radioactive tracer that can be incorporated in drinking water, or in the case of fish studies, in the tank water. It has been successfully used in hu‐ mans and other mammals for the study of hepatic intermediary metabolism in both normal and pathological conditions. It rapidly equilibrates with total body water and is distributed evenly into all tissues. It is a practical tracer for both short and long-term metabolic studies.

H2O is ideally suited for studying fish metabolism since it can be added to the tank water for an indefinite period, during which time it is incorporated into hepatic metabolites such as glycogen and glucose by specific enzymatic reactions in their biosynthetic pathways, as previously described for mammals. Applying these principles to free-swimming fish pro‐ vides an authentic metabolic profile that is unadulterated by anesthesia or infusion proce‐

H) is a stable isotope of hydrogen with a nucleus containing one proton and

H. The inherent sensitivity of 2

H *vs.* C-1

H counterpart, allowing tracer levels of 2

H because of kinetic isotope effects. The strength of a chemical bond

H-enriched metabolites compared to their tracees resulting in

H for any compound. Since metabolite transformation is governed in part

H. Metabolism of 2

H nucleus contains no neutron). In the NMR experiment, 2

between two atoms is dependent in part on their relative masses, hence a C-2

apparently slower rates of transformation. Moreover, with bulk levels of 2

**3. Deuterated water as a tracer of endogenous glucose and glycogen**

H2O has recently been used to study protein synthesis ki‐

Advances and Applications of Tracer Measurements of Carbohydrate Metabolism in Fish

H-enrichment from 2

H2O that are well

http://dx.doi.org/10.5772/54053

257

H resonates at a

H (at constant field and

H makes the bonds harder to

H transformation. This can discrim‐

H to be ob‐

H is not exact‐

H tracers, notably

H bond is

nitely maintained is possible. 2

will discuss its use in more detail.

hence the underlying principles of plasma glucose 2

dures that characterize the administration of classical carbon tracers.

with an equivalent number of nuclei) is about 0.9% that of 1

by breaking and formation of C-H bonds, the presence of 2

break thereby potentially slowing the rate of C-2

**synthesis**

**3.1. Overview**

**3.2. Basic principles**

one neutron (the 1

ly equivalent to that of 1

inate the transformation of 2

stronger than a C-1

different frequency compared to its 1

served in the presence of the tracee 1

Deuterium (2

( 2

2

#### **2.4. Measurement of glucose Ra by bolus injection and by primed infusion**

There are two approaches for quantifying plasma glucose Ra by isotope dilution. The first involves an intravenous injection of a tracer bolus and monitoring the decrease in specific activity or enrichment of plasma glucose as the tracer is being diluted by unla‐ beled glucose [75]. The dilution kinetics are best represented by more than one exponen‐ tial decay function indicating the presence of separate pools of glucose in the body with different clearance characteristics. Because of the relative simplicity of a single injection tracer delivery, this method has been applied in many fish species including kelpbass *Paralabrax clathratus* [73], seabass *D. labrax* [74], *Hoplias malabaricus* [76], common carp *C. carpio* [23]. However, this approach requires frequent blood sampling over a sustained period to adequately describe the complex tracer clearance kinetics. Limited by the num‐ ber of blood samples that could be drawn from the fish, the study described in [73] ex‐ trapolated the clearance kinetics from a smaller set of initial measurements, but the uncertainties of this approach were acknowledged. The alternative primed-infusion meth‐ od establishes a constant level of tracer in the bloodstream following an appropriate pri‐ ming dose. Under these conditions, the dilution of the tracer, measured from the ratio of specific activity or enrichment of infused label to that of blood glucose, is equal to the ratio of endogenous glucose appearance and tracer infusion rates. If glucose Ra is varia‐ ble (for example during meal ingestion) then the rate of infusion needs to be adjusted accordingly in order to maintain a constant ratio of infused glucose to blood glucose specific activity or enrichment. Compared to the bolus injection method, primed infu‐ sion measurements require fewer blood samplings; in fact the sole rationale for multiple blood samplings is to verify a constant ratio of infused to blood glucose specific activity or enrichment. Calculation of glucose Ra from steady-state isotope data is more robust compared to single injection since it is independent of the complex and often poorly de‐ fined clearance kinetics. However, a primed infusion requires catheterization of a vein to deliver the tracer over an extended period and is technically more difficult than a single bolus injection. While these procedures now allowed glucose Ra to be well determined, and by combination with dietary tracers can determine the contributions of absorbed and endogenous glucose production to glucose Ra, they do not inform the sources of en‐ dogenous glucose production. Novel methodologies of resolving the sources of glucose Ra using deuterated water (2 H2O) have been developed and can be integrated with primed-infusion glucose Ra measurements [80, 81, 82]. 2 H2O is ideally suited for fish met‐ abolic studies since it can be incorporated into aquarium water for an indefinite period and is rapidly incorporated into the fish body water such that the 2 H-enrichment level of the fish tissue water is also fixed for the duration in the 2 H-enriched tank water. As a re‐ sult of enzyme-catalyzed exchange reactions between bulk water and metabolite hydro‐ gens, many important precursor metabolites of glucose and glycogen biosynthesis, such as lactate, pyruvate and alanine also become rapidly enriched to the same level as body water for the study duration. Thus, a constant precursor enrichment that can be indefi‐ nitely maintained is possible. 2 H2O has recently been used to study protein synthesis ki‐ netics in the catfish *Ictalarus punctatus* [83] and gilthead seabream *S. aurata* [84] as well as blood glucose and hepatic glycogen turnover in seabass *D. labrax* [43, 85]. Because of its enormous potential for informing endogenous carbohydrate metabolism in fish, we will discuss its use in more detail.
