THE PARTITIONING OF ENERGY FROM DIGESTIBLE CARBOHYDRATES BY RAINBOW TROUT (ONCORHYNCHUS MYKISS)

A Thesis

Presented to

The Faculty of Graduate Studies

of

The University of Guelph

by

DOMINIQUE BUREAU

In partial fulfilment of requirements

for the degree of

Doctor of Philosophy

April, 1997

© Dominique Bureau, 1997

ABSTRACT

THE PARTITIONING OF ENERGY FROM DIGESTIBLE CARBOHYDRATES BY RAINBOW TROUT (ONCORHYNCHUS MYKISS).

Dominique Bureau Advisors :University of Guelph, 1997 Dr. J.B. Kirkland

Dr. C.Y. Cho

A series of studies was carried out to examine the partitioning of the energy from

dietary carbohydrates by rainbow trout. A pair-feeding technique, used for all the

experiments, consisted of feeding the same amount of a basal diet, low in digestible

carbohydrate, to the fish with different quantities of supplemental (digestible)

carbohydrate. One group of fish was fed the basal diet, alone, while two other groups

received supplemental glucose or gelatinized starch.

In the first study, glucose and gelatinized potato starch were the supplemental

carbohydrate fed to the fish. The energy from both carbohydrates was not efficiently

retained by the fish and did not affect nitrogen retention. A large proportion of the

energy from dietary carbohydrate could not be directly accounted for when comparing

intake of energy with the sum of fecal and non-fecal (nitrogenous) energy losses and

energy recovered in the carcass.

The second study was conducted to determine if the energy unaccounted for

could be due to catabolism and dissipation of the energy as heat. The fish were fed two

levels of supplemental gelatinized starch. Respirometry (oxygen consumption) was used

to estimate heat production by the fish. Feeding supplemental starch did not result in any

increase in heat production by the fish compared to that of fish fed the basal diet, alone.

An energy budget based on results from comparative carcass analysis and oxygen

consumption accounted for more than 90% of the intake of energy of the fish receiving

only the basal diet, whereas this proportion decreased for fish fed the basal diet with

supplemental starch.

In the third study, urinary glucose excretion of glucose was measured in non-

catheterized fish. This study showed that fish receiving supplemental carbohydrate

excreted a significant amount of glucose in their urine. This excretion represented a

significant portion (30%) of the unaccounted for carbohydrate energy losses in fish fed

high dietary levels of digestible carbohydrate. A large proportion of the supplemental

carbohydrate energy still needs to be accounted for.

The results from these studies indicate that dietary digestible carbohydrates (in

excess of what was provided by the basal diet, i.e. approximately 14%) contributes little

in terms of net energy to rainbow trout.

ACKNOWLEDGMENTS

The love and support from my parents, Denis and Henriette, have given me the

confidence to accomplish what I wanted to accomplish in life. I am forever grateful to

them. I wish my father was still here with us until this day. I also wish to thank my two

brothers, Germain and Frédéric, their spouses, Marie-Claire and Julie, and my niece,

Heidi, who helped me in the pursuit of this goal.

This work would not be achieved without the guidance from several peoples. I

would like to express my utmost gratitude to Dr. C.Y. Cho, who has been a mentor and a

friend to me. I am also very grateful to Dr. J.B. Kirkland for being an excellent advisor.

I appreciated his help, suggestions and friendship. Many thanks to the members of my

advisory committee, Drs. J.L. Atkinson, L.E. Nagy and W.D. Woodward, whose

comments, advices and encouragements were always helpful. Thanks to Drs. C.B.

Cowey and S.J Kaushik for their excellent suggestions and for teaching me some of the

fundamentals of fish nutrition.

I would like to express my gratitude to everyone who has helped me in any way

since I came to Guelph. I wish to thank the people from the Fish Nutrition Research

Laboratory, namely Greg Arndt, Andrew Harris, Ursula Wehkamp, Jantien van de

Grootest, Reginald Wade, Greg Page, Danielle McDonald, Pakinee Limwattanagura,

Karen Hayworth, Jennifer Cuts, and Paula Azevedo for their help and friendship. I

i

would also like to thank my friends from Dr. J.B. Kirkland's laboratory and the

Department of Human Biology and Nutritional Sciences as well as my friends from

hockey. I have appreciated the warmth of my Thai friends who always cared and cooked

delicious Thai food. Finally but not last, I wish to thank Vanida Khumnirdpetch for her

love and companionship, as well as her help and support during this undertaking.

ii

TABLES OF CONTENTS

Chapter ONE. INTRODUCTION....................................................................................1

Chapter TWO.LITERATURE REVIEW..........................................................................3

2.1 DIETARY CARBOHYDRATES..............................................................3

2.2 METABOLIC, PHYSIOLOGICAL AND NUTRITIONAL ASPECTS OF

CARBOHYDRATE UTILIZATION........................................................6

2.2.1 GLYCOLYSIS..................................................................6

2.2.2 TRICARBOXYLIC ACID CYCLE.................................10

2.2.3 OTHER METABOLIC PATHWAYS.............................11

2.2.4 METABOLIC AND PHYSIOLOGICAL ASPECTS OF

CARBOHYDRATE UTILIZATION IN FISH.................15

2.2.4.1 METABOLIC "BOTTLENECK" IN THE UTILIZATION

OF CARBOHYDRATE.......................................16

2.2.4.3 COMPLEX VS SIMPLE CARBOHYDRATES..............20

2.3 PARTITIONING OF DIETARY ENERGY IN FISH WITH

PARTICULAR REFERENCE TO CARBOHYDRATES.......................21

2.3.1 DIETARY SOURCES OF ENERGY - INTAKE OF ENERGY

(IE).............................................................................................. 25

2.3.2 FECAL ENERGY (FE)...............................................................26

iii

2.3.3 NON-FECAL ENERGY (UE+ZE)..............................................30

2.3.3.1 NITROGENOUS PRODUCTS........................................31

2.3.3.2 URINARY EXCRETION OF GLUCOSE.......................33

2.3.4 HEAT PRODUCTION................................................................38

2.3.4.1 BASAL METABOLIC RATE (HeE)...................40

2.3.4.2 HEAT INCREMENT OF FEEDING (HiE).........44

2.3.5 GROWTH AND RECOVERED ENERGY (RE)........................47

2.4 STATEMENT OF OBJECTIVES...........................................................51

Chapter THREE. COMPARISON OF THE UTILIZATION OF GLUCOSE AND

GELATINIZED POTATO STARCH BY RAINBOW TROUT

(ONCORHYNCHUS MYKISS) USING A COMPARATIVE CARCASS

ANALYSIS APPROACH................................................................................... 54

3.1 INTRODUCTION................................................................................... 54

3.2 MATERIALS AND METHODS............................................................59

3.3 RESULTS............................................................................................... 69

3.4 DISCUSSION......................................................................................... 78

Chapter FOUR: PARTITIONING OF ENERGY OF CARBOHYDRATE BY

RAINBOW TROUT (ONCORHYNCHUS MYKISS) : COMPARATIVE

CARCASS ANALYSIS AND RESPIROMETRY APPROACH........................82

4.1 INTRODUCTION................................................................................... 82

iv

4.2 MATERIALS AND METHODS............................................................84

4.3 RESULTS............................................................................................... 99

4.4 DISCUSSION.......................................................................................118

Chapter FIVE. URINARY EXCRETION OF GLUCOSE BY RAINBOW TROUT AT

INCREASING DIGESTIBLE STARCH INTAKES.........................................125

5.1 INTRODUCTION................................................................................. 125

5.2 MATERIALS AND METHODS...........................................................126

5.3 RESULTS AND DISCUSSION............................................................131

Chapter SIX. GENERAL DISCUSSION....................................................................144

REFERENCES............................................................................................................. 151

v

LIST OF TABLES

Table 3.1 Formulation of the experimental diets.....................................................61

Table 3.2 Analysis of the experimental diets...........................................................62

Table 3.3 Calculated nutrient intakes per fish in the main feeding trial over the 9-

week experimental period........................................................................64

Table 3.4 Apparent digestibility coefficients (ADC) of experimental diets calculated

from a pooled sample of two collection periods......................................72

Table 3.5 Performances and carcass characteristics of the fish fed the experimental

diets........................................................................................................ 73

Table 3.6 Estimation of the energy expenditure of the fish fed the experimental diets

................................................................................................................ 76

Table 3.7 Calculated energy values of the experimental diets and supplemental

carbohydrates.......................................................................................... 77

Table 4.1 Formulation of the experimental diets.....................................................88

Table 4.2 Chemical composition of the experimental diets.....................................89

Table 4.3 Experimental conditions for respirometry...............................................95

Table 4.4 Calculated nutrient intakes per fish over a 12-week experimental period

.............................................................................................................. 100

Table 4.5 Apparent digestibility coefficients (ADC) of experimental diets...........101

Table 4.6 Performance and carcass characteristics of the fish fed the experimental

vi

diets over a 12-week period...................................................................105

Table 4.7 Whole body carcass composition of the fish fed the experimental diets.107

Table 4.8 Energy budget of the fish fed the experimental diets for a 12-week period

in the main feeding trial........................................................................108

Table 4.9 Weight, nitrogen, and energy losses and calculated fasting heat losses of

rainbow trout during fasting trial...........................................................110

Table 4.10 Oxygen consumption and calculated heat production of fish measured in

the respirometry trial on the 3rd day of food deprivation (FHP) or

attributed to heat increment of feeding (HiE)........................................112

Table 4.11 Estimated heat losses per fish for 12-week main feeding trial...............116

Table 4.12 Calculated energy values of the experimental diets...............................117

Table 5.1. Urine flow rate (UFR) of fish fed the various experimental diets..........133

Table 5.2 Plasma and urine glucose concentrations and urinary glucose excretion of

the fish fed the experimental diets.........................................................134

vii

LIST OF FIGURES

Figure 2.1 Some pathways of glucose utilization in fish.............................................8

Figure 2.2 Scheme illustrating of the partitioning of dietary energy by fish according

to the convention of the NRC (1981)..................................................................23

Figure 2.3 Renal glucose reabsorption and excretion as a function of plasma glucose

in mammals........................................................................................................ 35

Figure 3.1 Growth curve of rainbow trout fed the experimental diets for a 9-week

period 74

Figure 4.1 Schematic representation of the experimental schedule...........................86

Figure 4.2 Growth curve of the fish fed the experimental diets for 12 weeks.........103

Figure 4.3 Example of the measured oxygen consumption of fish..........................113

Figure 5.1 Urinary glucose concentration (mM) of rainbow trout fed the various

experimental diets as a function of plasma glucose concentration.....................136

Figure 5.2 Estimated (calculated) urinary glucose excretion (mmol/kg per h) of

rainbow trout fed the various experimental diets as a function of plasma glucose

concentration.................................................................................................... 138

viii

LIST OF ABBREVIATIONS

ADC Apparent digestibility coefficientADP Adenosine diphosphateATP Adenosine triphosphateBW Body weightCHO CarbohydrateCP Crude protein (defined here as N x 6.25)d DayDE Digestible energyDGC Daily growth coefficientDM Dry matterDN Digestible nitrogenDP Digestible proteinDP/DEDigestible protein to digestible energy ratiodpm Disintegrations per minuteFBW Final body weightFE Fecal energy lossesFHP Fasting heat productionG Glucoseg GrammeG-6-P Glucose-6-phosphateGE Gross energyGFR Glomerular filtration rateGH Growth hormoneGTP Guanosine triphosphate3H Tritiumh HourHE Heat production, heat lossesHeE Basal metabolic rate, basal heat lossesHcE Heat of thermoregulation of body temperatureHdE Heat of digestion and absorption processesHjE Heat of voluntary activityHiE Heat increment of feedingHrE Heat of retention and interconversion of substratesHSD Tukey's honestly significant difference (minimum significant

difference)HSI Hepatosomatic indexHwE Heat of formation and excretion of waste products

ix

IE Intake of gross energyIBW Initial body weightIGF-1 Insulin-like growth factor 1IU International unitsJ Joulekg kilogrammekJ kilojouleL LitreLBW Live body weightMBW Metabolic body weightME Metabolizable energymg Milligrammesmin MinuteMJ MegajoulemM MillimolarN NitrogenNAD Nicotinamide adenine dinucleotide (oxidized form)NADHNicotinamide adenine dinucleotide (reduced form)NADP Nicotinamide adenine dinucleotide phosphate (oxidized form)NADPH Nicotinamide adenine dinucleotide phosphate (reduced form)NE Net energyNFE Nitrogen free extractPEG-4000 Polyethylene glycol (M.W. 4000)PFK PhosphofructokinasePK Pyruvate kinaseQ10 Temperature coefficientRE Recovered (retained) energyRN Recovered (retained) nitrogenRQ Respiratory quotientrpm Rotations per minuteSDA Specific dynamic actionSEM Standard error of the mean (pooled)TCA Tricarboxylic acid cycleTGC Thermal growth coefficientUE Urinary energy lossUFR Urine flow rateZE Branchial (gill) energy loss

Nota: This thesis is written in the styles and forms of the Journal of

x

Nutrition.The notation applied to energy terms is that suggested by NRC (1981).

xi

Chapter ONE. INTRODUCTIONChapter ONE. INTRODUCTION

Carbohydrates are, in general, the most economical source of food energy and

they compose the bulk of the diet of most monogastric animals. About one-half to three-

quarters of the digestible energy fed to pigs and poultry is supplied by starch and sugars.

Fish are poikilothermic (cold-blooded) animals and generally have a lower energy

expenditure than homeothermic (warm-blooded) animals because of the high basal

metabolic rate of the latter. Fish require less energy, but similar protein intakes, to

achieve the same growth as monogastric homeothermic animals, such as poultry, and

have a high ability to use protein as an energy source. Most fish also have a high

capacity to use dietary lipids as an energy source (Watanabe 1982). Replacing the more

expensive protein or lipid by the cheaper carbohydrate in fish diets has been the objective

of many studies in the past decades. Several studies have shown that salmonids fed diets

with high levels of dietary carbohydrates (e.g. 20-30%) exhibited relatively poor growth,

increased liver size, poor feed efficiency, poor energy retention and elevated levels of

blood glucose for extended periods of time (Philips et al. 1948, Luquet et al. 1975,

Edwards et al. 1977, Furuichi and Yone 1980, Hilton and Atkinson 1982, March et al.

1985, Beamish et al. 1986, Hilton et al. 1987, Higgs et al. 1992, Mazur et al. 1992).

Other researchers have concluded that 25-30% digestible carbohydrate in the diet of

rainbow trout had beneficial effects on protein and energy utilization (Pieper and Pfeffer

1

1980a&b, Kaushik et al. 1989, Kim and Kaushik 1992, Brauge 1994). The optimum

level and nutritive value of carbohydrate for fish is still a controversial subject (Pfeffer

1995).

Despite the very large number of publications on the subject, very few detailed

studies on the utilization and fate of energy from carbohydrate have been carried out. As

a result, the contribution of carbohydrate to the total energy requirement of fish still

remains unclear (NRC 1993). The study of data from feeding trials (e.g Hilton et al.

1987) shows that, at high levels of carbohydrate intake, a large portion of the energy

absorbed by the fish seems unaccountable for when comparing intake of energy (IE) and

recovered energy (RE), fecal energy losses (FE) and predicted heat losses (HE). There

have been very few attempts to directly or indirectly measure critical components of the

energy budget, such as heat increment of feeding (HiE) and urinary and branchial energy

losses (UE+ZE) of fish fed high levels of carbohydrates.

To determine how the energy from carbohydrate is partitioned, it is necessary to

build a detailed energy budget by quantifying its various components. It is necessary to

integrate several approaches, including apparent digestibility measurement, comparative

carcass analysis (comparative slaughter technique), respirometry (oxygen consumption),

and measurement of the urinary energy losses. A detailed energy budget could yield

much needed information regarding the metabolizable and net energy contents (ME, NE

respectively) of dietary carbohydrate and shed light on previous experimental results

2

which are sometimes contradictory.

3

Chapter TWO. LITERATURE REVIEWChapter TWO. LITERATURE

REVIEW

2.1 DIETARY CARBOHYDRATES.1 DIETARY CARBOHYDRATES

Carbohydrates are a very heterogeneous group with a basic chemical structure

composed of carbon, hydrogen and oxygen. As indicated by their name, they are

hydrated carbon atoms, whose empirical chemical formula can be expressed as C(H20)n

(Candy 1980). There are, however, many carbohydrates which do not fit exactly this

formula. Carbohydrates are, in general, the most economical source of food energy.

They compose the bulk of the diet of most domestic animals. The major function of

carbohydrate is as a fuel, to be oxidized and provide energy for other metabolic

processes. The most common forms in animal nutrition are mono, oligo and

polysaccharides made up of glucose molecules alone or in association with other

monosaccharides, such as galactose and fructose. Carbohydrates vary widely in their

potential nutritive value for animals. In cellulose, for example, the glucose units are

assembled via a 1-4 linkage. The enzyme cellulase affects the hydrolysis of these bonds

to release single glucose units. This enzyme is, however, absent in animals. It is found

in a number of other types of organisms (some bacteria and fungi) and most animals rely

on these organisms to digest and/or convert cellulose into utilizable nutrients, either in

the rumen or in the hindgut. Starch is a polymer of glucose, but the glucose units are

linked via 1-4 and 1-6 linkages. These bonds are hydrolyzed by enzymes, such as

4

amylase and maltase, which are widely distributed in animals.

Starch is generally the most abundant type of carbohydrate in the diet of

monogastric animals. Starch occurs in a granular form in the organs of plants. The word

starch may be derived from the Anglo-Saxon word stearc and has the connotation of

strength and stiffness, as applied to a fabric or to paper (Swinkels 1985). Starch granules

are deposited in the seeds, tubers, roots and stem of plants, as a reserve of food supply

for periods of dormancy, germination and growth. Starch is composed of tiny granules,

ranging from about 2 to 100 m diameter (Swinkels 1985). The granules are formed in

the plant after the deposition of a minute amount of insoluble polysacharrides, which

serves as a nucleus (hilum) around which further starch deposition occurs. As the

granules grow they often become elongated or flattened. The starch molecular chain

grows in an orientation perpendicular to the growing surface of the starch granule.

Whenever linear segments of the starch molecules are parallel to one another, hydrogen

bonding forces pull the chains together into associated crystalline bundles or micelles.

Starch granules contain crystalline bundles and amorphous regions (Swinkels 1985).

Most starches contain two types of glucose polymers: (1) a linear-chain molecule

termed amylose and (2) a branched polymer of glucose termed amylopectin. The two

fractions occur in differing proportion in starch from various botanical origins. Amylose

comprises 15-30% of the common starches (Swinkels 1985). Glycogen, the polymer of

glucose synthesized by animals, has the same chemical structure as amylopectin with a

slightly higher branching density (Candy 1980)

5

Gelatinization of starch

Starch granules in their native form are insoluble in cold water. This insolubility

is due to hydrogen bonds, formed either directly via neighbouring alcoholic OH groups

of individual starch molecules or indirectly via water bridges. The hydrogen bonding

forces are weak, but the large number of hydrogen bonds does not allow dissolution of

the starch granule in cold water. When starch granules are heated in water to

progressively higher temperatures, hydration in the amorphous regions of the starch

granules increases. This disrupts the weak hydrogen bonds between starch molecules and

causes swelling. This swelling disrupts the orderly radial orientation of the crystalline

bundles and the starch granules expand to become a greatly swollen reticulated network,

still held together by persistent crystalline bundles. The swelling is irreversible and this

process is refered to as gelatinization (Swinkels 1985). The impact of gelatinization on

the nutritive value of starch for fish will be discussed later.

6

2.2 METABOLIC, PHYSIOLOGICAL AND NUTRITIONAL ASPECTS OF

CARBOHYDRATE UTILIZATION.2 METABOLIC,

PHYSIOLOGICAL AND NUTRITIONAL ASPECTS OF

CARBOHYDRATE UTILIZATION

The percularities of the intermediary metabolism and utilization of carbohydrate

by fish have been reviewed extensively by Furuichi (1988), Cowey and Walton (1989),

and Wilson (1994). A scheme presenting some of the pathways of glucose utilization in

fish is presented in Figure 2.1.

2.2.1 GLYCOLYSIS.2.1 GLYCOLYSIS

All the enzymes of the Embden-Meyerhof pathway (glycolysis), well-defined in

mammals (Harper 1969), are present in teleost fish (Cowey and Watson 1989).

Glycolysis is the major route of glucose catabolism in fish tissues. This series of

reactions converts glucose to acetyl-CoA with the generation of high energy molecules,

such as ATP. In aerobic cells, glycolysis provides substrate for the TCA cycle in

mitochondria. Under anaerobic conditions, as in the white muscle of fish, pyruvate is

converted to lactate to regenerate NAD+, but this limits the amount of energy liberated

per mole of glucose catabolized.

7

Glycolytic activity is greatest per g of tissues in cardiac and skeletal muscle of

fish and lowest in tissues such as liver and kidney (Cowey and Walton 1989). A range of

published values for glycolytic enzymes in tissues of rainbow trout is provided by

Walton and Cowey (1982).

Glucose enters the glycolytic pathway by phosphorylation to glucose-6-phosphate

(G-6-P) through the action of the hexokinase enzymes (several isomers). The reaction is

accompanied by considerable loss of free energy as heat. Hexokinase has a high affinity

(low Km) for its substrate, glucose. It functions in mammals to supply glucose for

tissues, even in the presence of low blood glucose concentrations. In mammals, a

hexokinase with a high Km, glucokinase, has the function of removing glucose from the

blood following a meal. The glucose phosphorylating capacity was found to be several

times lower in fish than in mammals. Hexokinase is present in a lower level than any

other glycolytic enzyme in the liver of rainbow trout (Cowey and Walton 1989). The

hexokinase activity in the liver of trout (0.1-0.5 mol/min/g of wet tissue at 15C; Cowey

et al 1977a) is 5 to 20 times lower than that found in rat liver (2.5 mol/ min/ g of wet

tissue at 37C; Newsholme and Start, 1973).

8

Figure 2.1 Some pathways of glucose utilization in fish.

Legend is as follows: 1. hexokinase, 2. glucose-6-phosphatase, 3. phosphoglucomutase,

4. UDP-pyrophosphorylase, 5. glycogen synthetase, 6. glycogen phosphorylase, 7.

phosphoglucose isomerase, 8. phosphofructokinase, 9. fructose diphosphatase, 10.

aldolase, 11. triose-phosphate isomerase, 12. triose phosphate dehydrogenase, 13. 3-

phosphoglyceratekinase, 14. 2,3-phosphoglycerate mutase, 15. enolase, 16. pyruvate

kinase, 17. pyruvate carbokinase, 18. phosphoenolpyruvate carboxykinase, 19. lactate

dehydrogenase, 20. pyruvate dehydrogenase, 21. glucose-6-P dehydrogenase, 22.

gluconolactonase, 23. 6-phosphoglucobate dehydrogenase, 24. ribulose-5-P isomerase,

ribulose-5-P 3-epimerase, 25. transketolase, 26. transaldolase, 27. transketolase.

9

10

2.2.2 TRICARBOXYLIC ACID CYCLE.2.2 TRICARBOXYLIC ACID

CYCLE

Under normal aerobic conditions the pyruvate formed in the glycolytic

breakdown of glucose is not reduced to lactate, but is converted to acetyl-CoA, which is

oxidized to CO2 and water by the tricarboxylic acid (TCA) cycle. The presence of all the

enzymes of the TCA cycle (with the exception of succinyl thiokinase) was demonstrated

in rainbow trout and it has been shown that the TCA cycle functioned in a manner very

similar to that of mammals (Cowey and Walton 1989). The overall equation describing

glycolysis with subsequent oxidation of acetyl-CoA by the citric acid cycle is:

Glucose + 38 Pi + 38 ADP + 6 O2 6 CO2 + 38 ATP + 44 H2O

Equation 2.1

When 1 mol of glucose is combusted in a calorimeter to CO2 and water,

approximately 2870 kJ are liberated as heat. When oxidation occurs in the tissues, some

of the energy is not lost immediately as heat but is captured in high-energy phosphate

bonds. Under aerobic conditions, glucose is completely oxidized to CO2, water and 38

high-energy phosphate bonds are generated per molecule. The total energy captured in

ATP per mole of glucose oxidized is 1398 kJ, or approximately 49% of the energy of

combustion. Under anaerobic conditions, which prevail in the white muscle of fish,

pyruvate is converted to lactate with the net production of two high-energy phosphate

11

bonds per molecule of glucose. The total energy capture is about 74 kJ which represents

only 3% of the energy of combustion. Consequently, to provide a given amount of ATP

energy, about 20-fold more glucose must undergo glycolysis under anaerobic as

compared with aerobic conditions.

2.2.3 OTHER METABOLIC PATHWAYS2.2.3 OTHER METABOLIC

PATHWAYS

While the bulk of glucose may be metabolized through the glycolytic and the

TCA pathways to generate ATP, minor pathways exist that seem primarily designed to

provide certain tissues with specific molecules. G-6-P is an important compound, being

at the junction of several metabolic pathways (glycolysis, gluconeogenesis, hexose

monophosphate shunt, glycogenesis, and glycogenolysis).

Hexose monophosphate shunt

The hexose monophosphate shunt (also referred to as the pentose phosphate

pathway) is one of the most significant alternate pathways of utilization of glucose. The

major functions of the hexose monophosphate shunt are (1) the provision of reduced

12

NADP (NADPH) required by anabolic processes outside the mitochondria, such as the

synthesis of fatty acids and steroids, (2) the synthesis of ribose for nucleotide and nucleic

acid synthesis and 3) metabolism of reactive peroxides (hydrogen peroxide and fatty acid

hydroperoxides).

Glycogenesis and glycogenolysis

Glycogen synthesis and breakdown in the liver appear to be similar in fish and

mammals. Synthesis and breakdown are catalyzed by glycogen synthetase and glycogen

phosphorylase, respectively. Both enzymes occur in two forms, an a form with high

activity and a less active b form. These forms are interconvertible by phosphorylation or

dephosphorylation (Harper 1969, Cowey and Walton 1989). Glycogen synthesis and

breakdown play a key role in the maintenance of glucose homeostasis in mammals. In

fish, blood glucose does not appear to be tighly controlled and gluconeogenesis appears

to meet most of the glucose demand. Glycogenesis is quantitatively important in fish fed

high dietary levels of digestible carbohydrate (Hilton and Atkinson 1982). Glycogen

might be a significant source of energy for the fish under specific, energy-demanding and

acute conditions, such as during adaptation to seawater (Soengas et al. 1992).

Gluconeogenesis

13

In the absence of a dietary source of glucose this molecule must be formed from

non-carbohydrate precursors (Cowey and Walton 1989). The primary pathway involved

is that leading to pyruvate. Seven of the enzymatic steps of glycolysis are reversible and

so take part in gluconeogenesis. There are three non-reversible steps and alternate

enzymatic steps can by-pass them. Two molecules of pyruvate are required for the

formation of one molecule of glucose. This involves expenditure of six high-energy

bonds, four from ATP and two from GTP. It also requires a reducing equivalent in the

form of NADH for glyceraldehyde-3-phosphate formation. Since the conversion of

glucose into pyruvate by glycolysis only generates 2 ATP, the synthesis of glucose from

pyruvate is a process with a significant cost.

The main sites of gluconeogenesis in fish are the liver and the kidney (Cowey and

Walton 1989). The roles of gluconeogenesis are believed to be recycling of muscle

lactate, production of glycerol for lipogenesis, and synthesis of glucose from dietary

amino acids to supplement dietary intake. This last role may be essential to cover the

need of specific tissues, such as the brain and blood cells.

Gluconeogenesis in rainbow trout liver was shown to proceed less rapidly than in

omnivorous mammals (Cowey et al. 1977a) and this is in line with the low glucose

requirement of fish. Unpublished experimental evidence from the Fish Nutrition

Research Laboratory of the University of Guelph and another research institute (S.J.

14

Kaushik, personal communication) show that salmonids receiving diets with little (<8%)

or no digestible carbohydrate maintain remarkably stable blood glucose levels (2 - 3

mM) over the whole day (24h). While blood glucose levels may not be as tightly

controlled as in mammals on the upper level (i.e. unlike in mammals, the blood glucose

level in fish is allowed to remain high for an extended period of time), a minimum level

of blood glucose appears to be homeostatically maintained by fish, presumably to ensure

the supply of glucose to certain tissues, such as the brain and other nervous tissues, and

the blood cells, as in other animals (Cowey et al. 1977b).

Lipogenesis

The pathways of lipogenesis in fish are qualitatively similar to those in other

vertebrates. The liver is the main site of lipid synthesis in fish and is a far more

significant site of synthesis than the adipose tissues (Lin et al. 1977c). Hepatic fatty acid

synthetase of fish resembles that of mammals, but it appears that de novo fatty acid

synthesis is much less rapid in fish than in mammals (Sargent et al. 1989). Fatty acid

synthesis was shown to be affected by short-term and long-term nutritional factors, such

as the level of lipid or carbohydrate in the diet in salmonids and other fish species (Lin et

al. 1977b, Sargent et al., 1989). In salmonids, variation in the dietary lipid level appears

to be a more effective modulator of fatty acid synthesis than the level of dietary

carbohydrate (Sargent et al. 1989). It appears that lipid levels in excess of 9-10% of the

15

diet inhibit de novo fatty acid synthesis in salmonids (Brauge 1994).

2.2.4 METABOLIC AND PHYSIOLOGICAL ASPECTS OF CARBOHYDRATE

UTILIZATION IN FISH.2.4 METABOLIC AND PHYSIOLOGICAL

ASPECTS OF CARBOHYDRATE UTILIZATION IN FISH

Many feeding trials and metabolic studies have been carried out to ascertain the

extent of utilization of dietary carbohydrates by fish. The poor ability of fish to utilize

carbohydrate was illustrated early in the history of fish nutrition (Philips et al. 1948).

Glucose tolerance tests confirmed the limited ability of salmonids and other fish to

metabolize digestible carbohydrate. Oral administration of glucose (1 g) to yearling

rainbow trout (live body weight not given, Palmer and Ryan 1972) led to a linear

increase of blood glucose concentration over a 7h period. Post-prandial hyperglycemia

reached grossly elevated values of blood glucose exceeding 28 mM (500 mg/dL). Fish

metabolize glucose in a manner similar to the mouse but at a much slower rate. Plaice

metabolized 23% of injected [U-14C] glucose marker to CO2 after 14h (Cowey et al.

1975), rainbow trout 22-29% after 12h (Brauge 1994), and channel catfish 49% after 24h

(Saad 1989 cited by Wilson 1994). In the mouse, 32% of injected 14C from glucose is

expired as CO2 in 1h and 83% after 8h (Vrba 1966). The portion of carbon retained as

protein, fat or glycogen was approximately 20% in plaice after 14h and 16% in channel

catfish after 24h. The rate of glucose utilization (glucose replacement rate) was found to

16

be between 0.35-0.43 mg/min per kg body weight in coho salmon (Lin et al. 1977a)

which is much lower than rates observed in homeothermic animals (3 - 16 mg/min per kg

body weight). Glucose transit time (the average sojourn time of a glucose molecule in

the glucose mass) was 400 min in coho salmon compared to 30 - 100 min in

homeothermic mammals (Lin et al. 1977a).

Peripheral tissue glucose uptake does not appear to respond to increased blood

glucose levels in fish (Cowey et al. 1977b). From experimental observations reviewed

by Mommsen and Plisetskaya (1990), it appears that an insulin surge is present after a

meal rich in carbohydrates. This surge leads to increased binding of insulin to receptors,

as typically observed in mammals. However, glucose uptake by fish peripheral tissue

seems to be refractory to insulin probably due to a lack of the appropriate "machinery" to

utilize large amounts of glucose, e.g. insulin recruitable glucose transporters (Glut4),

poor rate of phosphorylation, or restricted entry of metabolites of carbohydrate for

oxidation through the TCA cycle.

2.2.4.1 METABOLIC "BOTTLENECK" IN THE UTILIZATION OF

CARBOHYDRATE.2.4.1 METABOLIC BOTTLENECK IN THE

UTILIZATION OF CARBOHYDRATE

17

The low glucose phosphorylating capacity in the tissues of fish has been

suggested as the most significant factor that could explain the poor ability of fish to

utilize carbohydrate (Cowey and Walton 1989). This view is supported by observations

by Tung and Shiau (1991) who found that the activity of hexokinase in the liver of

Tilapia (Oreochromis nilotica x O. aureus) was about half (in terms of NADH hydrolysis

under optimal conditions) the phosphofructokinase (PFK) activity. On the other hand,

experimental evidence from Furuichi (1988), who presented comparative data on the

activities of hexokinase and phosphofructokinase in yellowtail (carnivorous; optimal

dietary glucose = 10%), red sea bream (carnivorous; optimal dietary glucose = 20%) and

common carp (omnivorous; optimal dietary glucose = 30-40%), do not support this

hypothesis. In the study presented by Furuichi (1988), hexokinase had a similar activity

among species, but the activity of PFK varied according to their apparent ability to utilize

carbohydrate.

Other experimental evidence does not support the hypothesis of the limiting

hexokinase. Hilton and Atkinson (1982) observed that hepatic glucose-6-phosphate

dehydrogenase activity (G6PDH) increased progressively when dietary glucose

(substituting fish oil and cellulose) increased from 0 to 21% (50, 60, 97, 124, 201 mol

substrate utilized/min per kg wet body weight for diets with 0, 7, 11, 14 and 21% glucose

respectively). This progressive increase may indicate that amount of substrate (G-6-P)

for this enzyme is increasing with increasing digestible carbohydrate intake. This is

partly supported by observations of Atkinson et al. (1984a) who found that feeding a diet

18

with 30% glucose significantly increased fasting hepatic G-6-P and F-6-P levels of

rainbow trout compared to a diet without glucose. An increase in G6PDH activity in

response to an increase in digestible carbohydrate intake (and decrease in dietary lipid

intake) has also been observed by other investigators with other fish species (Lin et al.

1977b, Shimeno et al. 1993).

In most experiments involving feeding fish with increasing dietary levels of

digestible carbohydrates, a parallel increase in the relative size of the liver

(hepatosomatic index, HSI %) and in the deposition of glycogen is seen (Hilton and

Atkinson 1982, Kim 1989, Brauge 1994). This may indicate that overall carbohydrate

entering glycolysis, by phosphorylation through hexokinase, may exceed what can be

utilized and that the extra G-6-P is channelled toward the production of glycogen. This

is again discordant with the belief that hexokinase, and not another enzyme further down

the glycolytic pathway, is the bottleneck in the utilization of dietary carbohydrate (Hilton

and Atkinson 1982). An alternative explanation could be that hexokinase is actually the

limiting enzyme for the entry of glucose into glycolysis in the liver and other organs

(kidney, red muscle), but that in these organs, most of the glycogen is not directly

synthetized from glucose (e.g. glucose G-6-P G-1-P UDP-glucose glycogen;

Figure 2.1). In mammals, it has been suggested that a significant amount of dietary

glucose could be converted into gluconeogenic precursors in extrahepatic tissues before

being converted to glycogen (McGarry et al. 1987). Lactate and pyruvate are

particularily good substrates for glycogenesis (McGarry et al. 1987). In mammals,

19

minimal glycogenesis occurs in perfused liver or isolated hepatocytes when glucose is

used as sole exogenous substrate. However, when infused into rats, [1- 14C] glucose was

readily converted to liver glycogen, but with extensive randomization of the label of the

carbon atoms of the glucose contained in the glycogen. Randomization can only occur if

the glucose is first converted to trioses and subsequently resynthesized into glucose.

Randomization of 26-37% of the carbon of glucose contained in glycogen after injection

of [6 - 14C] glucose has been shown in fish (Nagai and Ikeda 1971). In mammals, it

appears that much of the lactate taken up by the liver is of splanchic origin (McGarry et

al. 1987). A significant amount of lactate could be derived from extrahepatic metabolism

of glucose and gluconeogenic amino acids in fish. Exogenous glucose is readily oxidized

by rainbow trout enterocytes as are glutamate and glutamine. Harris (1993) showed that

the hexokinase activity of the enterocyte is 8-fold that of the hepatocyte in rainbow trout

and that most of the carbon from catabolized glucose was found in lactate, pyruvate and

alanine. The formation of lactate, pyruvate and alanine by the trout intestine following

glucose uptake may play a role in the synthesis of liver glycogen.

The limited use of carbohydrate might be partially explained by a limited ability

of the carbohydrate metabolites to compete for entry in the TCA cycle. In vitro studies

on freshly isolated fish hepatocytes showed that alanine was preferentially oxidized to

CO2, whereas lactate was preferentially converted to glucose, even though both substrates

must pass through the pyruvate pool. The compartmentation of the pyruvate pool of the

cell and a mechanism that controls NADH level have been suggested as potential

20

explanations for the difference for the fate of the carbon skeleton of these two products

(Mommsen 1986, Suarez and Mommsen 1987).

The observed decrease in phosphoenolpyruvate carboxykinase (PEPCK) activity

(Hilton and Atkinson 1982), and increase in fructose diphosphatase (Hilton and Atkinson

1982) and PFK (Furuichi 1988) and G-6-PDH (Hilton and Atkinson 1982, Shimeno et

al. 1993) activities suggest that there is an increase in substrate circulation (cycling?)

through parts of the glycolytic-gluconeogenic pathways and the hexose monophosphate

shunt in response to increasing digestible carbohydrate intake.

2.2.4.3 COMPLEX VS SIMPLE CARBOHYDRATES.2.4.3 COMPLEX

VS SIMPLE CARBOHYDRATES

It has been observed in a limited number of occasions that high dietary levels of

digestible starch were more efficiently used than similar levels of dietary glucose by

teleost fish (Furuichi and Yone 1980, Wilson and Poe 1987, Tung and Shiau 1991, Shiau

and Lin 1993). The slower liberation of glucose from the starch via enzyme hydrolysis

leading to a slower appearance of the glucose in the circulation, compared to simple

glucose, might not overload the limited ability of the trout to utilize this molecule

(Buddington and Hilton 1987). Similarly, it has been observed that increasing the

21

feeding frequency of a diet high in digestible carbohydrate improved the utilization of the

glucose (Tung and Shiau 1991, Hung and Storebakken 1994), probably for the same

reasons.

22

2.3 PARTITIONING OF DIETARY ENERGY IN FISH WITH PARTICULAR

REFERENCE TO CARBOHYDRATES.3 PARTITIONING OF

DIETARY ENERGY IN FISH WITH PARTICULAR REFERENCE TO

CARBOHYDRATES

The catabolism of food is organized within the animal to harness energy and

substrates for use in anabolic and other life sustaining processes. The physiological

mechanisms which achieve this are very complex, allowing the catabolism of a large

variety of food molecules using the finite number of enzyme systems which are found in

animal tissues (Krebs and Kornberg 1957). One cannot directly measure the free energy

changes which occur in animals as the chemical energy of the diet is used to support life

processes. However, the dietary energy is ultimately either voided as feces and

metabolic wastes, dissipated as heat, or deposited in tissues. The law of conservation of

energy and the law of initial and final states are the fundamental principles that form the

basis of bioenergetics. Thus, if an increase of energy is found in one place, for example

in the body of an animal, an equal quantity of energy has been removed from another

place, i.e. the consumed food (NRC 1981).

Measurements of the energy cost of growth, particularly in livestock, are of great

practical importance, and the methodology and results have been summarized by Kleiber

(1975). Studies on energy utilization and expenditure of fish in captivity as well as

methodological approaches to the study of nutritional energetics in fish have been

23

extensively reviewed by Cho et al. (1982), Cho and Kaushik (1990), and Kaushik and

Médale (1994).

The nutrient content of feed can be determined by chemical analysis. Such

analyses measure the total amounts of chemicals/nutrients in the feed. In most cases, the

value of the feed as a source of nutrients is less than that indicated by chemical analysis

because a portion of the nutrient content may be unavailable. This usually implies that a

nutrient is present in the feed, in a form (e.g., in combination with some other

component) which resists the digestive processes of the fish. In other cases the nutrient

may be absorbed, but remain unused by the fish, because it cannot enter into the

metabolic processes of fish in its complexed form or because some of the nutrients are in

excess in relation to others. Such compounds are usually excreted through the gills or in

the urine. However, since there is a multiplicity of physiological processes involved in

the utilization of dietary fuels, an elaborate scheme has been developed, and generally

accepted by convention, to describe the losses which occur in the utilization of dietary

energy for tissue growth in animals. Measurements of these energy losses involve the

use of animals since this aspect of assessing nutritive value of a feed is a biological

evaluation rather than a physical or chemical analysis (Blaxter 1967). This convention is

described along with a comprehensive glossary by the NRC (1981) and is illustrated in

Figure 2.2.

24

Figure 2.2 Scheme illustrating of the partitioning of dietary energy by fish according

to the convention of the NRC (1981).

25

26

2.3.1 DIETARY SOURCES OF ENERGY - INTAKE OF ENERGY (IE).3.1

DIETARY SOURCES OF ENERGY - INTAKE OF ENERGY (IE)

Energy is not a nutrient. It is rather an end-product of absorbed energy-yielding

nutrients when they are oxidized and metabolized. Energy is an abstraction that can only

be measured in its transformation from one form to another (NRC 1981). All organic

compounds in fish feed release heat upon combustion, and thus are potential sources of

energy. The gross energy content of a food is measured by combustion, usually in a

heavily walled metal container (bomb) in an atmosphere of compressed oxygen. The

method is referred to as bomb calorimetry. Under these conditions the carbon and

hydrogen are fully oxidized to carbon dioxide and water, as they are in vivo. However,

the nitrogen is converted to oxides which is not the case in vivo. The oxides of nitrogen

interact with water to produce strong acids which can be estimated by titration, allowing

a correction to be applied for the difference between combustion in an atmosphere of

oxygen and catabolism in vivo. The gross energy intake of an animal is simply the

product of the quantity of food consumed and its heat of combustion (GE).

The gross energy value of the feed depends upon its chemical composition, the

mean values of heat of combustion of carbohydrate, protein and fat being 17.2, 23.6 and

39.5 kJ/g respectively (Brafield and Llewellyn 1982). Digestion, absorption and

utilization of the carbohydrates, lipids and proteins derived from the food are associated

with various biological losses of undigested and unutilized nutrients which would yield

27

energy if catabolized. Thus the measurement of the dietary energy value needs to be

assessed by both chemical and biological assays.

For salmonid fishes, lipids and proteins provide the primary dietary sources of

energy (Cho and Kaushik 1990). Dietary carbohydrates do not seem to play such an

important role. Physiologically, lipids and protein form an important part of the structure

of a fish, but the need for chemical energy can preclude their incorporation into the

tissues and may involve their catabolism as a source of energy. Thus, utilization of the

energy and nutrients of each diet depends both upon the level of intake and upon the

composition of the diet.

2.3.2 FECAL ENERGY (FE).3.2 FECAL ENERGY (FE)

The first task in evaluating the potential nutritive value of any feedstuff for

inclusion in a diet is the measurement of its digestibility. It is difficult to separate fish

feces from the water and to avoid contamination of the feces by the uneaten feed. This

problem has necessitated the use of very different approaches from those used to measure

digestibility for mammals and birds. This topic has been reviewed by Cho et al. (1982).

Before the feed components can serve as fuels for fish, they must be digested and

28

absorbed from the digestive tract. Some feed components resist digestion and a large

portion of these passes through the digestive tract to be voided as feces. The energy

which would have been liberated by the combustion of the fecal material is lost to the

animal and is referred to as the fecal energy loss (FE). The difference between the gross

energy of the food, and the gross energy of the feces derived from a unit quantity of

consumed food, is termed the digestible energy value. The digestible energy value of a

well-digested food would approach its gross energy value.

Feces are composed of the undigested food components and the unreabsorbed

residues of body origin. These residues are the remains of mucosal cells, digestive

enzymes and other secretions released into the digestive tract by the animal, together

with the residues of the microflora which inhabit the digestive tract. The residues not

arising directly from ingested food but from the metabolic activities of the animal are

referred to as metabolic residues. The heat of combustion of these materials represents a

loss of energy due to the process of digestion which is not derived from the food. This

energy loss is designated fecal energy of metabolic origin (FmE) and is influenced by the

characteristics of the food and the level of feed intake. Endogenous nitrogenous losses

have been estimated at around 2.7 - 3.3 mg/100 g live body weight per day or 123 - 144

mg/100 g dry diet consumed in common carp at 20C (Ogino et al. 1973). Endogenous

energy losses as protein (probably the most significant form of endogenous energy loss)

can, therefore, be estimated to be around 0.4 kJ/100 g live body weight per day or 20

kJ/100 g dry diet. This is small, being equivalent to less than 1% of the gross energy of

29

the diet. However, it does allow the description of "corrected" or "true" digestible energy

values which are greater than `apparent' digestible energy values. The term "true"

digestibility may be misleading since, to the animal, FmE losses are real and inevitable.

"Apparent" digestible energy (ADE) = IE - FE

"Corrected" digestible energy = IE - (FE - FmE)

Equation 2.2

Variation in the digestibility of foods is generally a major factor affecting the

variation in their usefulness as energy sources to the animal, since the fecal energy loss is

usually the major loss of ingested gross energy. Therefore, digestible energy values and

the digestibilities of individual nutrients such as protein, fat and carbohydrate should be

used to estimate levels of available nutrients in the feed ingredients for the formulation of

diets (Cho and Kaushik 1990). The commercially formulated diets normally used in fish

culture result in the loss to the animal of between 15 and 35% of the gross energy in the

feces. Fish, particularly salmonids, digest protein and fat very well. Fibre (cellulose,

hemicellulose, lignin) is indigestible by fish (Shiau 1989).

Digestibility of carbohydrates

Simple sugars are well absorbed by rainbow trout. However, starch in its native

30

form found in cereals and tuber, is poorly digested by rainbow trout (Cho and Slinger

1979, Bergot and Breque 1983). A summary of the digestibility of starch of various

origin is presented by Bergot (1993).

Buddington and Hilton (1987) observed that the intestine of rainbow trout did not

adapt to an increase of glucose or starch in the diet. The absorptive capacity for glucose

was good, but the activity of disaccharidases was depressed by a diet containing 25%

glucose. These authors hypothesized that enzymatic hydrolysis of carbohydrate could be

a regulatory step in the uptake of glucose by the intestine of trout.

Thermal treatment under moist conditions (hydrothermal treatment) leads to

swelling of the starch granule in a process refered to as gelatinization. It was shown in

numerous studies that gelatinization of starch by cooking or extrusion markedly increases

digestibility of starch in salmonids (Kim 1989, Pfeffer et al. 1990, Bergot 1993). It was

also observed that the apparent digestibility of starch varies inversely with the level in the

diet in salmonids (Bergot and Breque 1983, Kim 1989, Pfeffer 1995). Consequently, the

apparent digestibility coefficient (ADC) of complex carbohydrates is not additive

whereas the ADC of protein and lipid are (Kim 1989).

31

2.3.3 NON-FECAL ENERGY (UE+ZE).3.3 NON-FECAL ENERGY (UE+ZE)

Digestion of a diet leads to the absorption of amino acids, fatty acids and sugars

which are the principal metabolic fuels for the body. Catabolism of fat and carbohydrate

results in the formation of carbon dioxide and water, the fully oxidized products of

carbon and hydrogen respectively. However, the catabolism of amino acids yields

ammonia and that of purines and arginine yields urea (Kaushik and Fauconneau 1984),

which leads to the loss of combustible material by the fish. The loss of ammonia, urea

and other compounds (e.g. creatine, 3-methyl histidine) either through the gills (ZE) or

through the kidneys (UE) means that the digestible energy value of a diet overestimates

its fuel value to the fish. The physiologically available fuel value of the diet to the fish is

the metabolizable energy value defined as follows:

ME = IE - (FE + UE + ZE)

Equation 2.3

Determination of the metabolizable energy values of diets for fish is technically

difficult because of the need to quantitate both gill and urinary losses. However, Smith

(1971) attempted to overcome these difficulties and developed a procedure which

allowed the estimation of the metabolizable energy values of a number of feedstuffs

using rainbow trout. The method involves confinement of the fish and considerable

handling which are stressful to the fish. This may increase nitrogen loss (Hunn 1982)

32

and thus enhancing the loss of combustible matter. The resulting increase in nitrogen

output through the gills and kidneys, together with the low food intake attained by the

force-feeding of a single daily meal, might be expected to result in a negative nitrogen

balance and a low ratio of metabolizable to digestible energy values for many of the feed

ingredients studied. This suggests that energy losses via the gill and kidney measured in

these restrained fish were greater than would be the case for unrestrained fish feeding

normally.

2.3.3.1 NITROGENOUS PRODUCTS.3.3.1 NITROGENOUS

PRODUCTS

Catabolism of amino acids yields ammonia, in addition to carbon dioxide and

water. The pathway of degradation of purines yields another nitrogenous waste, urea.

The excretion of ammonia and urea leads to a loss of combustible material by the fish.

Most of these nitrogenous losses occur by excretion through the gills as ammonia

(Forster and Goldstein 1969), with some loss through the kidneys as ammonia and urea.

It has been determined that, in general, ammonia represents at least 85% of the

nitrogenous wastes whereas urea represent less than 15% (Kaushik and Cowey 1991).

The energy of combustion value of ammonia (82.3% N by weight) and urea (46.7% N by

weight) is 20.5 kJ/g (24.9 kJ/g N) and 10.5 kJ/g (22.5 kJ/g N) respectively (Brafield and

33

Llewellyn 1982). Since most nitrogen losses are as ammonia and the difference in the

amount of energy loss per g N between ammonia and urea is small, Cho and Kaushik

(1990) proposed that the loss of 1 g of nitrogen by fish under normal conditions could be

estimated as an energy loss of 24.9 kJ.

Protein sparing

The metabolizable energy value of a given feedstuff in a diet is not independent

of the composition of the diet, since it is the overall balance of the amino acids and

energy in the diet which influences the retention of protein by the body and hence

governs the loss of nitrogen products through the gills or in the urine.

As fish diets contain high levels of protein, a large amount of dietary energy is

provided as nitrogenous compounds that yield large quantities of nitrogenous catabolites.

Studies have indeed shown that increased provision of non-protein digestible energy can

considerably reduce the excretion of nitrogenous end products. The DP to DE ratio

(DP/DE) plays a very significant role in this phenomenon. Richly (1980) observed that a

decrease in dietary crude protein level (from 74% to 32%) with a concurrent increase in

the level of digestible carbohydrate from 9 to 53% led to a 45% decrease in nitrogen

excretory losses. In this study, however, feed efficiency was considerably affected by the

excess incorporation of carbohydrate in the diet. Kaushik and de Oliva-Teles (1985)

34

showed that an increase in digestible energy in the form of lipid or in the form of

carbohydrates led to a decrease in nitrogen excretion while improving energy retention.

Médale et al. (1991) observed that, at a given digestible protein level, an increase in DE

through incorporation of 30% gelatinized starch as opposed to raw starch in the diets of

rainbow trout led to a 10% reduction in ammonia nitrogen excretion. Other studies did

not observe any protein sparing effect of digestible carbohydrate (Hilton et al. 1987).

Protein sparing by digestible carbohydrate will only occur to the extent that carbohydrate

can be efficiently metabolized (i.e. catabolism of carbohydrate which allows the

harnessing of the chemical energy in high energy molecules and the fueling of anabolic

reactions).

Protein sparing has received a lot of attention in the past few years. It is current

practice to include very high lipid levels (30-35%) in fish feeds in an atttempt to spare as

much protein as possible. There is, however, a limit to protein sparing by lipid as

illustrated by the work of Cho et al. (1976). These authors showed that lipid in excess of

15% (in diet with 40% digestible protein) only exerted a very limited effect on protein

catabolism and heat increment of feeding.

2.3.3.2 URINARY EXCRETION OF GLUCOSE.3.3.2 URINARY

EXCRETION OF GLUCOSE

35

It has been suggested that fish fed an excessive amount of dietary glucose could

eliminate a portion of the glucose in the urine (Furuichi 1988, Shimeno 1991, Shiau and

Suen 1992). The excretion of a portion of the dietary glucose or of its metabolites in the

urine would mean that diets containing high levels of digestible carbohydrate would have

a ME content lower than that calculated only on the basis of nitrogeneous waste energy

excretion.

The dynamics of renal glucose reabsorption in teleost fish appear to be similar to

the those of mammals since these animals share the same type of renal glucose

reabsorption system that is a phlorizine-sensitive, sodium dependent and saturable

transport system (Dantzler 1989). A typical mammalian renal glucose reabsorption curve

as a function of plasma glucose and the pattern of the resulting glucose excretion is

presented in Figure 2.3.

Humans have a renal threshold concentration for excretion of glucose around 11

mM plasma glucose (Khoushanpour 1976). At plasma glucose levels below this

concentration, no glucose is lost into the urine. Progressively greater levels result in a

concomitant increase in urinary glucose excretion (curvilinear increase, also refered to as

"splay"). At 22 mM, the glucose reabsorption system is saturated and the urinary glucose

excretion increases at a rate parallel to the increase in plasma glucose (linear increase).

36

Figure 2.3 Renal glucose reabsorption and excretion as a function of plasma glucose

in mammals (adapted from Khoushanpour 1976).

37

38

The amount of information on the physiology of glucose tubular reabsorption in

the kidney of fish is relatively limited. There are, to the knowledge of this author, no

published studies on the renal threshold for glucose excretion in fish. Experimental

evidence as to the excretion of glucose by fish fed high dietary levels of digestible

carbohydrates remains mostly unpublished. Nonetheless, urinary excretion of glucose

has been observed in hyperglycemic fish. Yokote (1970) and Kakuta and Namba (1989)

observed glucosuria in hyperglycemic fish suffering from Sekoke Disease (caused by

peroxidized dietary lipids). French reseachers collected urine of rainbow trout fed diets

with 30% gelatinized starch and found that urinary excretion of glucose was negligible

(S.J. Kaushik, personal communication). The fish were fixed with a urinary bladder

canula, and urine was collected in a latex sleeve attached to the end of the canula. This

technique may be stressful and it is not known if the fish in that study maintained normal

feed intakes.

In the most detailed study on the topic, the incidence and degree of glucosuria

(described as negative, mild, moderate or severe) was measured in two carnivorous

fishes, red sea bream and yellowtail (Furuichi 1983). Yellowtail fed diets with 10 or

20% glucose had a high incidence and degree of glucosuria. The fish fed 20%

gelatinized potato starch had a low incidence and degree of glucosuria, while the fish fed

10% gelatinized starch did not show any glucose in their urine. In red sea bream,

glucosuria was highest in the fish fed a diet containing 25% glucose, intermediate in fish

fed diets with 25% dextrin and very low in fish fed 25% gelatinized starch. While only

39

qualitative descriptors of glucose concentration in the urine were provided, and blood

glucose concentration of the fish fed the various diets was not measured, this remains the

most informative study published to date. It demonstrates that urinary excretion of

glucose occurs in these species of fish and that it is determined by the source and level of

carbohydrate in the diet. Quantitative information on urinary excretion of glucose, along

with information on the relationship between blood glucose levels and urine glucose

excretion of fish, is needed, however, to determine the extent of the energy losses

involved.

The excretion of a portion of the dietary glucose or of its metabolites in the urine

would, therefore, mean that diets containing high levels of digestible carbohydrate would

have lower ME values than would be calculated only on the basis of nitrogenous waste

(ammonia and urea) energy excretion. Measurement of urinary excretion of glucose by

rainbow trout is essential to obtain a clearer picture of the partitioning of energy from

dietary carbohydrate by this species.

2.3.4 HEAT PRODUCTION.3.4 HEAT PRODUCTION (HE)

The nutrients absorbed from the digestive tract are either catabolized or stored as

new tissue components. Part of the energy released by catabolism of the nutrients is

40

ultimately released as heat, so that the energy balance can be determined either by

measuring heat production, or by estimating the change in body energy content from

weight and carcass composition changes (Blaxter 1989). The latter system is referred to

as body balance and requires observations over an appreciable portion of the animal's

growth phase.

Life proceeds in a thermodynamically unstable state and its continuation is

dependent on the balance between dietary energy intake and heat production as a

consequence of life-maintaining processes (Kleiber 1975). In fact, if the diet provides

less energy to the animal than is needed to sustain life processes and to support its

voluntary activities, body tissues will be catabolized in addition to the food.

Furthermore, animals also need energy to grow, reproduce, and support physical activity

(NRC 1981).

Heat is liberated by animals as a consequence of the transfer of the chemical

energy of nutrients to energy-rich molecules, such as ATP, during the metabolic

transformation of dietary substrates into biologically important substances, or during the

hydrolysis of ATP to do physical or chemical work (Kleiber 1975). The maintenance of

cellular homeostasis, the formation of new tissue or the turnover of existing tissues as

well as physical activity require the production, transformation and catabolism of a large

variety of molecules. However, these reactions do not harness all the energy, and a

significant portion of the energy is dissipated as heat at each step. The rate at which heat

41

is liberated is an indication of the intensity of ongoing reactions. This is designated

metabolic rate (Kleiber 1975). An important concept is that of basal metabolic rate

which, as the term implies, is the minimum rate of metabolic activity needed to sustain

the structure and function of the body tissues. The ingestion of food increases the

metabolic rate, as a consequence of the extra work due to the ingestion, digestion and

utilization of the food. This increase is termed the heat increment of feeding. Physical

activity also increases metabolic rate due to work done against internal and external

frictional forces. These three components of animal metabolism lead to the release of

energy as heat (HE) from the ME derived from the food. Clearly energy released as heat

is not available for weight gain (increase in body energy), which can only occur if the

dietary ME intake exceeds the rate of heat production. As might be anticipated, if the

ME intake is less than the rate of heat production, the difference will be provided by

catabolism of body tissues and weight loss will ensue. An intake of ME in excess of heat

production will be stored in the body as energy retained in new tissues (Cho and Kaushik

1990).

2.3.4.1 BASAL METABOLIC RATE (HeE).3.4.1 BASAL

METABOLIC RATE (HeE)

Fish require a continuous supply of energy for those functions of the body

42

immediately necessary for maintaining life, regardless of whether or not the animal is

consuming food. A fish deprived of food obtains this energy by catabolizing body

reserves of fat and protein. In fed fish, this requirement for maintenance energy is

supplied by the food, thus obviating the catabolism of body tissue. Among the

requirements for maintenance, a major portion of the energy is spent on basal metabolism

(HeE) and a smaller portion of energy is spent for voluntary or resting activity (HjE)

such as minor bodily movements (Cho and Kaushik 1990).

By definition, the basal metabolism is the minimum rate of energy expenditure

needed to keep the animal alive and this has highest priority in the maintenance

requirement. The basal metabolic rate is measured when the animal is in the

post-absorptive state, and is in a state of muscular repose at an environmental

temperature which is thermoneutral (Blaxter 1989). Definition of basal metabolic rate

for fish precludes the latter condition but makes it necessary to specify the temperature at

which the metabolic rate is measured.

Brett (1972) showed that the maintenance requirements of poikilothermic animals

were 1 to 3% of those of mammals which maintain high body temperatures (e.g. 37C).

It is more difficult to ensure that fish are in a state of muscular repose than in terrestrial

mammals because they need to maintain their orientation in the water and this entails

some muscular activity. Thus, basal metabolism can be measured by extrapolation to

zero activity from fish swimming at different rates (Smith 1989). Some fish, such as

43

rainbow trout, will spend considerable periods resting on the bottom of their tanks,

maintaining their position in quiet water with minimal activity. The energy expenditure

of fasting rainbow trout under these conditions can be regarded as a close approximation

to basal metabolism (Cho and Kaushik 1990).

Cho et al. (1976) found that fasting, resting rainbow trout at 15C released 59 to

63 kJ/kg per day as heat which is equivalent to 40 kJ/kg body weight to the 0.824 power

(BW0.824). These measurements were made with groups of fish whose individual weights

were in the range 96-145 g and used oxygen consumption to estimate the heat losses.

Kaushik and Gomes (1988) obtained similar results (46-60 kJ/kg per day at 18 C with

150 g rainbow trout).

Fasting heat production (FHP) was measured by Smith et al. (1978a) using direct

calorimetry. These authors found that FHP at 15C was 204 kJ/kg BW0.75 for 1-57 g

rainbow trout. This is a much higher estimate than that of Cho et al. (1976) and Kaushik

and Gomes (1988). On the other hand, this estimate is similar to the fasting heat

production rate of mammals (290 kJ/kg0.75; Kleiber 1975). The scientific value of the

FHP estimate of Smith et al. (1978a) is questionable, since it is in disagreement with the

widely recognized concept that fish, being poikilothermic animals, have a much lower

HeE than mammals. This apparent overestimation of the FHP of trout by Smith et al.

(1978a) may be attributable to the insensitivity of the method, direct calorimetry, when

used with aquatic poikilothermic animals, which release little heat and live in an aquatic

44

environment, which has high heat capacity. The reliability of other estimates of heat

losses, such as heat increment of feeding (Smith et al. 1978b), obtained by direct

calorimetry, is, therefore, also questionable.

The fasting heat production represents approximately 60-70% of the total heat

production (HE) of fish under normal cultured conditions. Variation in water

temperature has a great effect on the fasting heat production of fish. Cho and Slinger

(1980) measured the fasting heat production of rainbow trout (live weights from 47 to

136 g) at temperatures of 7.5, 10, and 15C. Water temperature had its largest effect on

fasting heat production between 7.5 and 10C, such an increase in temperature resulting

in a doubling of heat production. A further increase in water temperature to 15C

resulted in a 50% increase in heat production. Cho (1991) proposed the following

equation to calculate the HeE of salmonids as a function of the temperature (T) and their

bodyweight (BW):

HeE = (-1.04 + 3.26T - 0.05T2)*kg BW0.824

Equation 2.4

45

2.3.4.2 HEAT INCREMENT OF FEEDING (HiE).3.4.2 HEAT

INCREMENT OF FEEDING (HiE)

Ingestion of food by an animal which has been fasting results in an increase in the

animal's heat production. This expenditure of energy due to feeding is referred to by

several terms: heat increment of feeding (HiE), extra heat, specific dynamic action

(SDA), calorigenic effect, and diet induced thermogenesis (Cho and Kaushik 1990). The

factors contributing to the heat increment of feeding are considered to be: (1) formation

and excretion of metabolic wastes (HwE), (2) transformation and interconversion of the

substrates and their retention in tissues (HrE) and (3) the digestion and absorption

processes (HdE) (NRC 1981). Therefore, the net amount of energy available for

maintaining the life process and gaining growth is the net energy (NE) and this is

deducted for all losses caused by food intake:

NE = IE - FE - UE - ZE - HiE

where HiE = HwE + HrE + HdE

Equation 2.5

The main biochemical basis for heat increment is the energy required for the

ingested amino nitrogen to be deaminated and excreted (HwE) (Kleiber 1975). The

energy expenditures associated with food ingestion and digestion (HdE) are very small

compared to that associated with the metabolic work (HwE + HrE) (Brody 1945,

46

Emmans 1994). The physiological basis of this increased heat production includes the

post-absorptive processes related to ingested food, particularly protein-rich food and the

metabolic work required for the formation of excretory nitrogen products, as well as the

synthesis of proteins and fats in the tissues from the newly absorbed, food-derived

substrates such as amino acids and fatty acids (Blaxter 1989).

Cho and Kaushik (1990) estimated that the HiE associated with feeding a

maintenance ration is approximately one-third of the total HiE and the rest (two-thirds) is

the portion associated with the productive gain. Cho et al. (1976) observed that diets

containing 6% fat and either 30 or 47% of digestible protein resulted in similar rates of

oxygen consumption. However, increasing the fat level in the lower protein diet resulted

in a substantial reduction in oxygen consumption, which is the classical effect of lipid

upon HiE. Increasing the fat level in the higher protein diet had practically no effect

upon oxygen consumption, presumably because of the metabolic work associated with

the higher influx of amino acids provided by the high protein diet (Cho et al. 1982).

Carnivorous fish consume high protein diets and excrete much of the digested

nitrogen. The HiE, as a fraction of the digestible energy, varies with the level of protein

and fat in the diet. It is, however, remarkably independent of dietary composition when

expressed on the basis of nitrogen intake. It was concluded that protein metabolism is

the major factor contributing to the heat increment of feeding (Cho and Kaushik 1990,

Ross et al. 1992). Cho and Kaushik (1990) estimated the HiE of salmonids fed a

47

balanced diet to be approximately 30 kJ/g digestible N. Values for HiE for salmonids,

therefore, range from 8 to 12% of digestible energy intake.

Contribution of dietary carbohydrates to heat increment of feeding

It has been suggested that digestible carbohydrates are possibly also significant

contributors to HiE (Beamish et al. 1986, Médale et al. 1994). Beamish et al. (1986)

observed that oxygen consumption was greater in fish fed a diet rich in digestible

carbohydrate (35% crude protein, 10% lipid, 29% glucose) than in fish fed a diet rich in

lipids (35% crude protein, 22% lipid). The authors suggested that feeding digestible

carbohydrates increased the HiE. However, in that study, the fish fed the high

carbohydrate diet had lower growth rate and nitrogen retention. The greater catabolism

of protein could be responsible for the greater oxygen consumption of the fish fed the

high carbohydrate diet compared to the fish fed the high lipid diet. The results from that

study, in fact, closely resembles those of Cho et al. (1982) who reported a very

significant reduction in the oxygen consumption (reduction in heat increment) of fish fed

a 36% digestible protein diet when the lipid level was increased from 6% to 11 and 16%.

This suggests that the effect on HiE observed by Beamish et al. (1986) was in fact more

related to the variation in dietary lipid and the sparing of protein than to the digestible

carbohydrate itself.

48

In mammals, it has been shown that utilization of glucose as an energy source

only entails an HiE equal to approximately 1% of the GE of glucose. Converting glucose

into glycogen costs 5% of the energy of glucose as HiE whereas converting glucose into

lipids entails an increase of HiE equal to at least 20% of its gross energy (Blaxter 1989).

Information of this kind is very limited for fish. Because of the similarity in the

pathways of glucose utilization in mammals and fish, energy cost of glucose utilization

are expected to be similar in these types of animals.

2.3.5 GROWTH AND RECOVERED ENERGY (RE).3.5 GROWTH AND

RECOVERED ENERGY (RE)

ME which is not dissipated as heat is retained within the body as new tissue

elements. In growing animals, part of the recovered energy (RE) is stored as protein and

part as fat, but as the animal approaches its mature size an increasing proportion of the

RE is stored as fat; the relative importance of protein and fat deposition depends upon a

great number of factors in addition to the maturity of the animal (Cho and Kaushik

1990). The balance of the available amino acids of the dietary protein and the amount by

which the dietary energy intake exceeds the energy expended as heat are the two major

factors. Proteins of higher biological value promote greater protein deposition than those

49

of low value (Blaxter 1989, Kaushik and Cowey 1991). Marginal excesses of energy

intake over energy expended as heat result in the deposition of a larger proportion of the

RE as protein. As the excess energy intake increases, the total amount of protein

deposited increases, but the proportion of the energy retained as fat increases at an even

faster rate, so that increasing energy intake leads to an increase in the amount of energy

retained as fat (Cho and Kaushik 1990).

In almost all cases, retention of energy and the deposition of new tissue results in

an increase in the weight of the animal, and the weight gain of young fish is usually a

reliable indicator of the adequacy of the nutritional and management regimes.

Unfortunately, the rate of weight gain is not a quantitative measure of energy retention

for two reasons, firstly, because deposition of fat reduces the water content of the body,

thus changing the energy value per unit weight of the living animal, and secondly,

because the energy content of fat and protein per unit weight are so different (Blaxter

1989, Cho and Kaushik 1990). Fat is usually deposited in adipose tissue in association

with relatively little water resulting in a heat of combustion for fish adipose tissue of 31

kJ/g, whereas protein is usually deposited in such tissues as viscera and muscle in

association with a great deal of water resulting in a heat of combustion for fish muscle

tissue of 6 kJ/g (Cho and Kaushik 1990). This means that it is impossible to equate

weight gain with RE without a simultaneous estimate of body composition. Hence

comparisons of weight gain per unit of food consumed (feed efficiency) are only useful if

the energy value of the food and weight gain are known (Cho and Kaushik 1990).

50

This problem of interpreting productivity data is further complicated by the

different metabolic processes by which dietary energy is stored as either fat or protein

(Emmans 1994), the relatively simple synthesis of lipids being 74-90% efficient whereas

the much more complex synthesis of protein is only 35-60% efficient for rats, poultry,

swine and ruminants (Pullar and Webster 1977, Emmans 1994) and 40-50% for rainbow

trout (Cho and Watanabe 1987). For proper estimation of the energy efficiency of a diet,

it is, therefore, not only necessary to know the energy gain but also the composition of

the gain in terms of protein and lipids (Emmans 1994).

Contribution of dietary carbohydrate to recovered energy

Hilton et al. (1987) observed in the feeding of rainbow trout that crude corn

starch and glucose were poor contributors in terms of productive energy (energy stored

as protein and fat from the portion of the consumed ration which exceeds the quantity

used for maintenance purposes). Crude starch, at 25% of the diet, had a productive

energy value of 2.17 kJ, or 12.6% of its gross energy, while glucose had a productive

energy value of 3.99 kJ, or 24.6% of its gross energy. This denotes a poor ability of

these carbohydrates to contribute to the deposition of energy in the animal. Under

normal conditions, they probably do not contribute much to the sparing of protein, are

not efficiently converted into lipid, and energy stored as glycogen is quantitatively small.

51

This is in agreement with the results of Hilton and Atkinson (1982) who found very few

indices of lipogenesis from carbohydrates in rainbow trout fed increasing levels of

dextrose and decreasing levels of lipid.

Warmwater omnivorous fish, such as channel catfish, apparently have a relatively

high capacity to transform carbohydrates into lipid (Likimani and Wilson 1982). The

relative amount of lipid synthesized from carbohydrate in salmonids is, on the other

hand, still debated because of contradictory experimental evidence. Lin et al. (1977b)

found stimulation of fatty acid synthesis by feeding a high carbohydrate/low lipid diet

compared to a high lipid/low carbohydrate diet. Lipid deposition rates exceeding the rate

of digestible lipid intake have been observed in trout fed diets containing high levels of

gelatinized starch (Médale et al. 1994). Kaushik et al. (1989) observed very high

apparent lipid retention efficiency (160%) in rainbow trout fed a diet containing 30%

gelatinized corn starch, which was interpreted as indicating a very significant conversion

of carbohydrate into lipid. Brauge (1994) found incorporation of the radioactivity of

injected glucose into lipid in rainbow trout. However, only 2% of the total amount of

radioactivity injected was incorporated into lipid after twelve hours for a diet with a very

low lipid level. Conversion of glucose into lipid in fish fed a diet containing 10% lipid

was, however, depressed. It may be concluded that conversion of glucose into lipid is

probably low in salmonids fed a practical diet with moderate lipid levels.

Feeding high levels of digestible carbohydrate has been shown to greatly increase

52

liver size and the overall weight of viscera, mainly as a result of storage of glycogen, and

in some cases lipid, in the liver (Hilton and Atkinson 1982, Hilton et al. 1987, Brauge et

al. 1994). While a small, but non-negligible, fraction of the energy from digestible

carbohydrate may be retained as glycogen and lipids, these compounds are mainly stored

in the liver (Brauge 1994) and, therefore, may contribute very little by weight to the

productive or marketable portion of the animal, i.e. the dressed carcass. For studies on

the nutritive value of dietary carbohydrate, it is, therefore, not only necessary to look at

the live weight, energy, protein and lipid gains but also at the dressed carcass yield.

2.4 STATEMENT OF OBJECTIVES.4 STATEMENT OF OBJECTIVES

Very few detailed studies on the partitioning of the energy from carbohydrate

have been carried out, despite the enormous number of publications on various aspects of

carbohydrate metabolism and utilization in fish. As a result, the contribution of

carbohydrate to the total energy requirement of fish remains unclear and the subject of

debate. A study of data from available feeding trials (e.g. Hilton et al. 1987) shows that,

at high levels of carbohydrate intake, a large proportion of the energy absorbed by the

fish seems unaccounted for when comparing IE and RE, FE and predicted heat losses.

53

There have been very few attempts to directly or indirectly measure critical components

of the energy budget, such as HiE and UE+ZE of fish fed high levels of carbohydrates.

The principal objective of this thesis is, therefore, to determine how the energy

from carbohydrate is partitioned by rainbow trout (Oncorhynchus mykiss W.). This is an

attempt to answer the simple but unanswered question: Where is the energy from

digestible carbohydrate going in rainbow trout ? This thesis presents three studies with

different scopes and goals, but all centered around this principal objective.

A first study had the objective of establishing the energy budget of rainbow trout

at high intake of digestible carbohydrates using a dietary model and pair-feeding protocol

designed to specifically look the effects of carbohydrates. The energy budget of this

study is based on the digestibility and comparative carcass analysis approach described

by Cho and Kaushik (1990).

A second study had a wider scope, but used the same dietary model and pair-

feeding protocol as the first study. It integrated several measurements, including

digestibility, comparative carcass analysis, and respirometry (oxygen consumption) in an

attempt to build a more detailed energy budget for the trout. This more detailed budget

included estimates of heat losses, namely basal energy metabolism (HeE) and heat

increment of feeding (HiE), to determine if the apparently "missing" energy from

digestible carbohydrate is dissipated as heat.

54

Finally, a third study was designed to measure the extent of urinary glucose

excretion in an attempt to determine if urinary energy loss (UE) could explain the

apparent discrepencies when accounting for the energy budget of trout at a high intake of

digestible carbohydrates.

55

Chapter THREE. COMPARISON OF THE UTILIZATION OF GLUCOSE AND

GELATINIZED POTATO STARCH BY RAINBOW TROUT (ONCORHYNCHUS

MYKISS) USING A COMPARATIVE CARCASS ANALYSIS

APPROACH.Chapter THREE. COMPARISON OF THE UTILIZATION OF

GLUCOSE AND GELATINIZED POTATO STARCH BY RAINBOW TROUT

(ONCORHYNCHUS MYKISS) USING A COMPARATIVE CARCASS ANALYSIS

APPROACH.

3.1 INTRODUCTION.1 INTRODUCTION

It is widely recognized that carnivorous fish, such as salmonids, have a poor

ability to utilize dietary carbohydrates, but utilize dietary protein and lipid well. Fish fed

with diets rich in carbohydrate show, in general, poor growth and feed efficiency, and

elevated blood glucose for extended periods of time. The level of digestible

carbohydrate for carnivorous fish in the diet should be below 20% (Hilton and Atkinson

1982, Furuichi 1988). The effect of dietary carbohydrate on the growth of fish has been

the subject of several experiments, but very few actually examine the fate of the ingested

carbohydrate. The fate of the energy from carbohydrate remains obscure since fish, in

general, maintain a good feed intake on high carbohydrate diets but have relatively poor

feed efficiency and energy retention (Hilton et al. 1987, Kim 1989, Brauge 1994).

56

It has been suggested that complex but digestible carbohydrate, such as

gelatinized starch, could be utilized more efficiently than simple carbohydrates, such as

glucose, by fish (e.g. Tung and Shiau 1991). Glucose derived from gelatinized starch

may appear more slowly in the circulation and this might allow for a better metabolic

utilization of glucose (Buddington and Hilton 1987).

Choosing the appropriate dietary model

The formulation of the diets and the feeding protocol are probably the most

critical issues in the planning of a study to determine the utilization of dietary

carbohydrate energy. In animals, the major role for carbohydrate is to provide a supply

of fuel for various chemical reactions. Unlike their role in plants, carbohydrates only

have a minor structural role in vertebrate animals. They are catabolized or converted into

various molecules, such as glycerol, amino acids, fatty acids and nucleic acids, which

makes them difficult to follow without the use of sophisticated techniques, such as

radioisotopic tracers.

Several dietary models have been used to study the nutritional value of dietary

carbohydrate for fish. The dietary model and feeding system used may have a significant

impact on the estimate of the nutritive value of carbohydrate. Protein and lipid are used

as energy sources, and the utilization of dietary carbohydrate may be affected by the

57

levels of dietary protein and lipid present. The dietary levels (or levels of intake) of

carbohydrate may have a great impact on the estimate of the energy values of the diet,

since the response of fish to dietary carbohydrate is not always linear or additive in

nature (Hilton et al. 1987, Kim, 1989).

One of the most widely used dietary models to study the nutritive value of

carbohydrates has been the one in which increasing levels of digestible carbohydrates are

included in a diet series, while the protein and digestible energy contents are kept

constant by substituting carbohydrate for fish oil and cellulose (Hilton and Atkinson

1982, Higgs et al. 1992, Mazur et al. 1992). The fish are fed the diets to near-satiety or

at a predetermined level. A limitation of this method is that high levels of dietary

cellulose are known to affect performance of salmonids (Hilton et al. 1983). Cellulose is

indigestible to carnivorous fish. It decreases the gut transit time and may affect the

digestibility of certain nutrients (Shiau 1989). Hilton et al. (1983) showed that cellulose

in excess of 8% in the diet affected the growth performance of fish by limiting feed

intake. The main limitation of the method is that energy from carbohydrate substitutes

that of lipids. It is sometimes difficult to determine if changes observed in energy

expenditure can be attributed to increasing carbohydrate intake or to decreasing lipid

intake and this greatly complicates the analysis of energy expenditure.

A variation of this dietary model has been used by Kim (1989), Médale et al.

(1994) and Brauge (1994). It involves substituting either a digestible starch source

(gelatinized corn starch) for lipid and poorly digestible starch (raw corn starch). Diets

58

with high levels of digestible carbohydrate and low levels of lipids (e.g. 6%) are

compared with control diets which have low levels of digestible carbohydrate but contain

high levels of raw, poorly digestible, starch (20-38%) and relatively low levels of lipids

(5-15%). High levels of poorly digestible dietary starch may act as dietary fibre and may

limit the feed intake and growth performance of fish. The bulk of that type of diet is,

perhaps, shown by less than optimal feed efficiency values (Kim and Kaushik 1992).

Another widely used dietary model may be referred to as "protein-sparing"

(Bergot 1979 ; Pieper and Pfeffer 1980a). Increasing levels of carbohydrate replace part

of the protein in a diet series. Since amino acids are also used as an energy source, and

amino acid intakes are a determining factor in growth, the dietary level of protein

influences the utilization of the carbohydrate. This type of formulation also greatly

complicates the determination of the partitioning of energy from carbohydrate.

A main concern of the dietary models described above has been to equalize

nutrient or energy contents of the experimental diets. It has been suggested that diets

should be designed on the basis of ration allowances rather than feed formulation

(Moreau et al. 1995). Experimental diets should be regarded as the sum of different feed

ingredients, rather than a mix of ingredients. This concept is behind a rather unusual

dietary model used by March et al. (1985). These authors used diets containing various

levels of dextrose, protein and energy. The diets were fed at different rates to allow the

same protein intake. Pieper and Pfeffer (1980b) also used a dietary model that was

59

designed according to a similar idea. It involved a basal diet composed of fish meal,

casein and a vitamin premix which was either fed alone or as a mixture made of 7 parts

of basal diet and 3 parts of various sources of carbohydrate (sucrose, gelatinized starch

and lactose). The "diluted" diet was fed at 100% of a fixed feeding rate (1.5-2.0% LBW)

and the "undiluted" basal diet was fed at only 70% of these rates. The authors achieved

good results in controlling the nutrient intakes. The experimental design led to

inconclusive results, since only two groups of 11 fish were used per treatment and no

statistical analysis was performed. This dietary model, nonetheless, appears to be

appropriate for analyzing specifically the nutritional value (expressed as energy value) of

dietary carbohydrates. Protein, lipid, mineral and vitamin intakes are equalized thus

simplifying comparisons among diets. The elimination of masking factors should aid the

determination of the partitioning of energy from carbohydrate and the efficiency of the

utilization of carbohydrate.

Described herein is a feeding trial which attempted to determinate the utilization

and partitioning of supplemental glucose and gelatinized potato starch by rainbow trout.

A pair-feeding protocol similar to the one used by Pieper and Pfeffer (1980b) and the

digestibility - comparative carcass analysis approach proposed by Cho and Kaushik

(1990) were used in this trial.

60

3.2 MATERIALS AND METHODS.2 MATERIALS AND METHODS

Fish, Diets, and Feeding

Rainbow trout were obtained from a commercial fish farm (Spring Valley Trout

Farm, Petersburg, Ontario). The fish, weighing an average of 33 g, were stocked (45

fish/tank) in nine rectangular fibreglass tanks (30 L) supplied with a mixture of well

water and city water at a rate of about 3 L/min. Each tank represented one experimental

unit. The tanks were individually aerated, and water temperature was controlled

thermostatically at 15C. Photoperiod was maintained on a 12 h light : 12 h dark

schedule. The animals were treated in accordance with the guidelines of the Canadian

Council on Animal Care (CCAC 1984) and the University of Guelph Animal Care

Committee. Three dietary treatments were each allocated to three blocks of three tanks

using a complete block design.

The diet formulation (Table 3.1) involved the dilution of a high fish meal diet

(basal diet) with 40% glucose (Cerelose, Corn Products, Summit-Argo, Illinois) or 40%

gelatinized potato starch (Bind up, Sakamoto Feed, Japan). Fish meal and fish oil were

obtained from a local feed manufacturers (Martin Feed Mills, Elmira, ON). Gelatinized

potato starch was used because it is a highly digestible type of starch (Cho and Kaushik

1990). The chemical composition of the experimental diets is presented in Table 3.2.

Celite AW521 (Celite Corp., Lompoc, CA), a form of diatomaceous silica, was included

61

in the diet to serve as a digestibility indicator (acid-insoluble ash). The diets were mixed

using a Hobart mixer (Hobart Ltd, Don Mills, Ontario) and pelleted using a laboratory

steam pellet mill (California Pellet Mill Co., San Francisco, California). The feed pellets

were subsequently dried in a current of air at room temperature for 24h. The diets were

kept at 4C until used. The amount of diet needed weekly was then kept at room

temperature.

62

Table 3.1 Formulation of the experimental diets.

Diets1

Ingredients (g) Basal HG HS

Fish meal, herring, 68% CP 42.0 42.0 42.0

Fish oil 6.0 6.0 6.0

Celite AW5212 1.2 1.2 1.2

Vitamin premix3 0.6 0.6 0.6

Mineral premix4 0.6 0.6 0.6

Glucose 0.0 40.0 0.0

Starch, potato, gelatinized 9.6 9.6 49.6

Total (absolute weight) 60 g 100 g 100 g

1 Diet designation, Basal = basal diet, HG = 60% basal + 40% glucose, HS= 60% basal + 40% gelatinized potato starch2 Celite AW 521 (acid-washed diatomaceous silica) is a source of acid-insoluble ash3 provides per kg of basal diet: Retinyl acetate, 3750 IU; Cholecalciferol, 3600 IU; All-rac--tocopheryl acetate, 45 IU; Menadione sodium bisulfite, 15 mg; 1- Ascorbic acid, 600 mg; Cyanocobalamine, 0.0225 mg; d-biotin, 0.225 mg; Choline chloride, 1500 mg; Folic acid, 7.5 mg; myo-Inositol, 225 mg; Niacin, 225 mg; d-Calcium pantothenate, 45 mg; Pyridoxine.HCl, 15 mg; Riboflavin, 10.5 mg; Thiamin.HCl, 4.5 mg.

63

4 Provides per kg of basal diet: Sodium chloride, 3000 mg/kg; Ferrous sulfate, 63 mg/kg; Manganese sulfate, 86 mg/kg; Zinc sulfate, 144 mg/kg; Copper sulfate, 25 mg/kg; Potassium iodide, 8 mg/kg.

64

Table 3.2 Analysis of the experimental diets1.

Composition2 Basal HG HS

Dry matter, % 94.1 92.2 95.3

Crude protein, % 47.0 29.1 27.9

Digestible protein, % 44.3 27.2 25.5

Lipid, % 17.7 11.3 9.3

Ash, % 10.6 6.6 6.5

Nitrogen-free extract, % 18.8 45.2 51.6

Gross energy, MJ/kg 21.3 18.2 18.8

Digestible energy, MJ/kg 20.3 16.9 15.5

DP/DE, g/MJ 21.8 16.0 16.5

1 Diet designation, Basal = basal diet, HG = 60% basal + 40% glucose, HS= 60% basal + 40% gelatinized potato starch2 Dry matter, crude protein, ash according to standard AOAC (1995) methods. Lipid content determined to method of Bligh and Dyer (1959). Nitrogen-free extract obtained by difference. Gross energy by bomb calorimetry. Digestible protein = crude protein * apparent digestibility of protein (Table 3.4) and digestible energy = gross energy * apparent digestibility of gross energy (Table 3.4).

65

Feeding trial

The fish were fed the experimental diets for a period of 9 weeks. A pair-feeding

protocol was used to achieve equal consumption of the basal diet for all dietary

treatments but different intakes of carbohydrate. The high glucose diet (HG) was hand

fed to fish to near-satiety three times daily. The high starch (HS) and basal diet (Basal)

were then fed at levels equal to 100% and 60% respectively of the feed consumption of

the fish from the same block fed the HG diet in order for the fish in all three tanks in the

same block to receive the same amount of basal diet. Total feed, nutrient and energy

intakes of the fish from this study are presented in Table 3.3. Fish were weighed and

counted every 3 weeks. Five fish were sampled at the beginning and at the end of the

experiment for carcass analysis. Hepatosomatic index (HSI%) and relative weight of the

dressed carcass (whole carcass - viscera) of 3 fish per tank were also determined every 3

weeks. Daily growth coefficient (DGC%) was calculated according to Iwama and Tauz

(1981) and Cho and Woodward (1989) as follows:

Install Equation Editor and double-click here to view equation.

66

Table 3.3 Calculated nutrient intakes per fish in the main feeding trial over the 9-

week experimental period.

Diets1

Intakes Basal HG HS

Feed, g/fish 41.1 68.5 69.0

Dry matter, g/fish 38.7 63.2 65.8

Crude protein, g/fish 19.3 19.9 19.3

Digestible protein, g/fish 18.2 18.6 17.6

Lipid, g/fish 7.3 7.7 6.4

Ash, g/fish 4.4 4.5 4.5

Nitrogen free extract, g/fish 7.7 31.0 35.6

Gross energy, kJ/fish 904 1280 1338

Digestible energy, kJ/fish 860 1187 1100

1 Diet designation, Basal = basal diet, HG = 60% basal + 40% glucose, HS= 60% basal + 40% gelatinized potato starch

67

Digestibility trial

At the end of the feeding trial, the fish were moved to a different aquatic system

equipped with Guelph feces settling columns (CYAQ-2) to determine digestibility of the

diets according to the method of Cho et al. (1982). The aquatic system consisted of three

fibreglass tanks in each unit, which all drained through a common drainpipe, and a single

standpipe placed over an acrylic settling column (10 cm diameter x 40 cm high). The

tanks each measured 55 cm x 40 cm and had a sloped bottom. Each tank was loaded

with a maximum of 3 kg of fish. The velocity of the water flow was adjusted to

minimize settling of the feces in the drainpipe and maximize recovery of the feces in the

settling column.

The fish were acclimated to both the tanks and the dietary regime for more than

one week before collection of feces began. The fish were fed three meals daily between

0900 and 1600 hours, according to the same pair-feeding protocol as described earlier.

One hour after the last meal, the drainpipe and the settling column were brushed out to

remove feed residues and feces from the system. One-third of the water in the tanks was

drained to ensure that the cleaning procedure was complete. At 0830 hours the following

day, the settled feces and surrounding water were gently withdrawn from the base of the

settling column into a large centrifuge bottle. These feces were free of uneaten feed

particles and considered to be a representative sample of the feces produced throughout

the 24-hour period. Immediately after collection of the feces, the fish were fed again

68

according to the normal procedure, allowing repeated sampling over several days. The

feces were centrifuged at 4,000 x g for 20 min and the supernatant discarded. The feces

were then freeze-dried, ground and stored at -20C to await analysis.

Chemical analyses

Diets, feces, initial and final carcass samples were analyzed for crude protein

(%N x 6.25) using a Kjeltech auto-analyzer (Model # 1030, Tecator, Hoganas, Sweden)

and for gross energy using an oxygen bomb calorimeter (Model # 1341 EB, Parr

Instruments, Moline, Illinois). Digestion indicator (celite) was determined using the

acid-insoluble ash (AIA%) indicator method used by Atkinson et al. (1984b).

Energy budget

The following energy budget (NRC 1981) was used for this study to compare the

partitioning of dietary energy by the fish fed the experimental diets:

IE = FE + UE + ZE + RE + Balance

Equation 3.2

69

where :

IE = Gross energy intake

FE = Fecal energy (egested energy)

UE = Urinary energy

ZE = Branchial energy

RE = Recovered energy

Balance = Other unmeasured losses, including heat production

(HE) and errors from other measurements

IE, FE and RE were calculated based on the heat of combustion of the diet, feces

and carcass samples measured by bomb calorimetry. In this study, certain assumptions

were made, some of which will be examined in later chapters. For example, non-fecal

losses (UE + ZE) were calculated based on estimated soluble nitrogen losses made

according to Cho and Kaushik (1990) as follows:

[ Digestible Nitrogen intake (g) - Retained Nitrogen (g) ] * 24.9 kJ/g

Equation 3.3

The gross, digestible and metabolizable energy values of the diets and

supplemental carbohydrates were calculated according to NRC (1981). The energy

expenditure of the fish fed the basal diet alone served as baseline and the digestible,

metabolizable and net energy for gain (NEgain, productive energy) value of the

70

supplemental glucose or gelatinized starch were calculated from the difference in the

energy expenditure of the fish fed the basal diet with the supplemental carbohydrate and

the fish fed the basal diet alone as follows:

Statistical Analysis

Data were analyzed using the general linear model (GLM) using the SAS/STAT

software (SAS 1988) where appropriate using a complete random block design. Means

of dependent variables were compared using Tukey's honestly significant difference

(HSD) test, with an =0.05 (Steel and Torrie 1980). Pooled SEM and minimum

significant difference according to Tukey's HSD test were provided for each dependent

parameter.

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71

3.3 RESULTS3.3 RESULTS

Apparent digestibility coefficients for dry matter, crude protein and gross energy

are presented in Table 3.4. The apparent digestibility coefficients were calculated using

pooled samples from two collection periods because of the limited amount of sample

collected from the fish fed the basal diet alone. No statistical analyses could be

performed on these calculated apparent digestibility coefficients. The apparent

digestibility coefficient (ADC) of protein was high for all the diets which is reflective of

the good quality of fish meal used in these diets. The ADC of energy were high for the

Basal and HG diets, but lower for the HS diet. The ADC of energy of the various

experimental diets were in line with expected values, based on the ADC for protein of the

fish meal used in this study, and the ADC of energy of the various ingredients (fish meal,

fish oil, gelatinized potato starch) presented by Cho and Kaushik (1990).

Growth performances and carcass characteristics of the fish fed the experimental

diets are presented in Table 3.5 and the growth curves are shown in Figure 3.1. The final

body weights (FBW) of fish fed the experimental diets were not significantly different.

There was a significant reduction in dressed weight percentage of the fish receiving

supplemental glucose or starch. Dressed carcass weights were, however, not

72

significantly different. Feed efficiency of the fish fed the Basal diet was within the

expected range for this type of diet and size of fish (Cho 1992). Feed efficiency was

significantly diminished with supplemental carbohydrate allocation since no

improvement in growth was observed, despite greater physical amount of feed consumed.

Retained nitrogen (RN) was not significantly affected by carbohydrate supplementation.

The energy budget of the fish fed the experimental diets is presented in Table 3.6.

Feeding digestible carbohydrate resulted, in both cases, in a small increase in FE.

UE+ZE of the fish fed the HS diet was significantly lower than that of the fish fed the

Basal and HG diet. This is attributable to slightly lower digestible protein intake (but

similar crude protein intake) of the fish fed the HS diet. Minor differences in crude

protein, digestible protein and lipid contents among the experimental diets could be

attributed to variability in the analysis of these parameters. UE+ZE values were

calculated using expected nitrogen excretion as the only variable. There was no

difference in RE. The energy balance (which included HE) increased significantly with

the supplemental carbohydrate regardless of the source. Energy balance represented 52%

of the digestible energy fed for the fish fed the basal diet and 62 and 63% of the

digestible energy fed for the fish fed the HG and HS diets respectively. Overall, there

were no differences between glucose and pre-gelatinized potato starch with regard to

dietary energy utilization. The calculated GE, DE, ME and NE for gain (NE gain) values

of the supplemental glucose and gelatinized potato starch are presented in Table 3.7. The

HG and HS diets had low NEg values compared to that of the Basal diet. NEg values of

73

the supplemented carbohydrates were very low compared to their calculated DE and ME

values.

74

Table 3.4 Apparent digestibility coefficients (ADC) of experimental diets calculated

from a pooled sample of two collection periods.

Diets1

ADC Basal HG HS

Dry matter, % 86.3 86.1 86.6

Crude protein, % 94.2 93.3 91.3

Gross energy, kJ/g 95.1 92.7 82.2

1 Diet designation, Basal = basal diet, HG = 60% basal + 40% glucose, HS= 60% basal + 40% gelatinized potato starch

75

Table 3.5 Performances and carcass characteristics of the fish fed the experimental diets1.

Diets2

Parameters Basal HG HS SEM HSD3

Feed, g/fish 41.1 68.5 69.0 N/A4 N/A4

Final body weight, g/fish 85.0a 90.0a 90.0a 2.2 11.3

Feed efficiency (gain:feed) 1.23a 0.81b 0.81b 0.04 0.18

Daily growth coefficient, % 1.89a 2.03a 2.03a 0.06 0.23

HSI, % 1.17b 4.27a 4.63a 0.30 1.5

Dressed carcass, % 87.5a 84.2b 83.5b 0.76 3.8

Dressed carcass weight, g 74.4a 75.7a 75.2a 1.7 8.8

76

Retained nitrogen, g/fish 1.33a 1.34a 1.44a 0.05 0.26

1 Initial body weight = 33 g/fish2 Diet designation, Basal = basal diet, HG = 60% basal + 40% glucose, HS= 60% basal + 40% gelatinized potato starch3 Tukey's honestly significant difference. Means in a same row sharing a same letter are not significantly different. 4 Not applicable, independent variable

77

Figure 3.1 Growth curve of rainbow trout fed the experimental diets for a 9-week

period.

The diet designation is as follows: Basal = basal diet, HG = 60% basal + 40% glucose,

HS= 60% basal + 40% gelatinized potato starch.

78

79

Table 3.6 Estimation of the energy expenditure of the fish fed the experimental diets.

Components Diets1

(kJ/fish per 84 d) Basal HG HS SEM HSD

IE 904 1280 1338 N/A2 N/A2

FE 44c 93b 145a 1 7

UE+ZE3 42a 42a 36b 1 3

RE 371a 411a 409a 18 60

Balance4 447b 733a 749a 12 91

1 Diet designation, Basal = basal diet, HG = 60% basal + 40% glucose, HS= 60% basal + 40% gelatinized potato starch2 Not applicable, independent variable.3 Based on calculated nitrogen losses 4 Balance was obtained by difference (IE-FE-(UE+ZE)-RE) and will include heat losses (HE), as well as other unmeasured losses, such as

the possible loss of energy as glucose in the urine and through the gills, and the errors from the other components of the energy budget.

Table 3.7 Calculated energy values of the experimental diets and supplemental carbohydrates.

Feed Energy1 GE DE ME NEg

kJ/g kJ/g kJ/g kJ/g

Experimental diets2

Basal 21.3 20.3 19.4 16.1

HG 18.2 16.9 16.2 10.2

HS 18.8 16.8 16.4 10.8

Supplemental carbohydrates

Glucose 13.9 12.1 12.1 1.5

Gelatinized potato starch 15.1 11.6 11.6 1.4

1 GE= gross energy, DE= digestible energy, ME= metabolizable energy, NEg= net energy for gain2 Diet designation, Basal = basal diet, HG = 60% basal + 40% glucose, HS= 60% basal + 40% gelatinized potato starch

3.4 DISCUSSION.4 DISCUSSION

Feeding digestible carbohydrate energy as glucose or gelatinized starch resulted

in poor retention of the extra-energy (about 10% of the extra carbohydrate energy fed)

and only a small increase in fecal energy waste (FE). Such results are in accordance with

results of a previous study which showed that dietary glucose had a poor net energy value

for rainbow trout (Hilton et al., 1987). In contrast, the results from the present study

appear to be in contradiction with the conclusion drawn by Kim (1989) from a study on

the energy value of dietary carbohydrate. Kim (1989) concluded that gelatinized starch

was as effective a source of energy as lipid, on a digestible energy basis. Comparable

growth rates (DGC%) and energy and feed utilization efficiencies were obtained with the

HS diet used in the present study and a diet of similar composition (high digestible

carbohydrate) used by Kim (1989). This leads the present author to believe that the

difference between the conclusions from the present study and Kim (1989) may reside

simply in the interpretation of results or basis for comparison, i.e. the performance of the

fish fed the "control" diet. The lower digestible carbohydrate "control" diet used in the

study of Kim (1989) contained 40% digestible protein, 38% raw (poorly digestible)

starch and only 10% lipid. This diet was shown to support suboptimal growth

performance and poor nitrogen retention. A diet with a higher protein content and a

lower raw starch content was shown to sustain significantly higher growth than the low

digestible carbohydrate "control" diet and the high digestible carbohydrate diet in the

study of Kim (1989).

Feeding a complex carbohydrate, gelatinized potato starch, did not lead to higher

retention of energy or higher growth than a simple carbohydrate, glucose. This is in

contradiction with the suggestion of Buddington and Hilton (1987) and with the findings

of Tung and Shiau (1991) and Wilson and Poe (1987) who worked with tilapia and

catfish respectively. It is also partly in contradiction with the results of Hung and

Storebakken (1994) who observed an improvement in the efficiency of carbohydrate

energy utilization when diets rich in digestible carbohydrate were continously-fed instead

of meal-fed. This was apparently due to the fact that the low but continuous level of

glucose absorbed in continuously-fed fish did not overload the metabolic capacity of the

animal. The poor growth performance and feed efficiency obtained in that study does

not lend much credibility to the results.

It may be argued that gelatinized potato starch is highly digestible and that the

release and absorption of glucose from that type of starch is probably almost as rapid as

absorption of simple glucose. However, in a recent study, the present author observed

no difference in the efficiency of utilization of starches of various digestibilities (raw or

gelatinized starches of various botanical origins) incorporated at 20% in rainbow trout

diet (unpublished results).

The results from the present study suggest that (at similar intakes) there is no

advantage, in terms of energy utilization, to use a complex (digestible) source of

carbohydrate, such as gelatinized starch, compared to a simple source of carbohydrate,

such as glucose in salmonid diets. Based on personal observations by this author, the diet

containing 40% gelatinized potato starch appeared more palatable to the fish. The use of

gelatinized starch may, therefore, be more interesting from an experimental point of view

and may allow greater intake by the fish.

The "balance" component (unmeasured energy losses, such as heat losses and

errors of other components of the energy budget) for the fish fed the Basal diet was close

to expected heat losses calculated according to Cho and Kaushik (1990). On the other

hand, feeding the digestible carbohydrate resulted in a great increase in the "balance"

component. Possible explanations for this greatly inflated balance component of the fish

fed the carbohydrates could be an increase in heat loss by the fish or an increase in the

non-fecal losses (UE+ZE). Since, no direct or indirect measurement of heat production

of the fish was performed, it is impossible to to determine if the energy was really

dissipated as heat. Moreover, the estimate of non-fecal losses in this study was only

based on calculated (expected) losses of nitrogenous compounds (Equation 3.2). Some

hypothesis, including the existence of urinary glucose, need to be tested. If glucose or its

metabolites were to have been excreted in the urine, the non-fecal losses (UE+ZE)

component would be underestimated, and this would lead to an overestimation of the ME

content of the diet. The extent of glucose excretion via the urine in fish fed various

levels of dietary glucose from different dietary levels or sources of carbohydrate has

never been quantified and should receive some attention. Heat production and urinary

losses must be measured to describe more completetly the energy budget of rainbow trout

fed these high carbohydrate diets.

Chapter FOUR: PARTITIONING OF ENERGY OF CARBOHYDRATE BY

RAINBOW TROUT (ONCORHYNCHUS MYKISS) : COMPARATIVE CARCASS

ANALYSIS AND RESPIROMETRY APPROACHChapter FOUR

PARTITIONING OF ENERGY OF CARBOHYDRATE BY RAINBOW

TROUT (ONCORHYNCHUS MYKISS) COMPARATIVE CARCASS ANALYSIS

AND RESPIROMETRY APPROACH

4.1 INTRODUCTION4.1 INTRODUCTION

The effect of dietary carbohydrate on the growth of carnivorous fish has been the

subject of several experiments, but very few actually examined the partitioning of the

energy from dietary carbohydrates. It was reported in the previous chapter that fish fed

with high carbohydrate diets maintained good feed intake, but that the energy derived

from carbohydrate seemed mostly unaccounted for when comparing RE, FE, and UE+ZE

(nitrogenous compounds) of these fish. Two unaccounted sources of energy loss that

could be responsible for this "missing" energy could be an increase in heat production

(HE) or in urinary and branchial excretion of glucose.

Beamish et al. (1986) observed that fish fed a diet with a high glucose content

consumed more oxygen than fish fed a diet with the same protein level but rich in lipid.

They suggested that the increase in digestible carbohydrate intake could result in an

increase in heat increment of feeding (HiE). Other the hand, the fish fed the diet high in

glucose had a lower protein retention than the fish fed the diet high in lipid. The

catabolism of amino acids for energy may be responsible for the increase in HiE obtained

with the high glucose diet. There are few published studies looking specifically at the

effect of increasing the digestible carbohydrate intake on the heat production of fish.

This information could help determine if the unaccounted for energy loss of fish fed high

levels of digestible carbohydrate is due to an increase in HE.

While this study has a slightly wider scope than the one presented in the previous

chapter, it uses the same dietary model and pair-feeding protocol with minor

modifications. It integrates several types of measurements, including digestibility,

comparative carcass analysis, and measurement of oxygen consumption to estimate heat

production in an attempt to build a more detailed energy budget for the fish. This energy

budget includes an estimate of heat losses, namely basal energy metabolism (HeE) and

heat increment of feeding (HiE), to determine if the apparently missing energy from

carbohydrate is dissipated as heat.

4.2 MATERIALS AND METHODS4.2 MATERIALS AND METHODS

Experimental design

Three feeding trials were conducted in parallel for this study. These trials began

on the same day and each used twelve groups of 100 rainbow trout (initial live body

weight = 8.1 g/fish). The trials are identified throughout this chapter as feeding, fasting

and respirometry trials. Figure 4.1 provides a schematic representation of the

experimental design and experimental schedule.

The feeding trial was conducted over a 12-week period. The fish were fed

according to the dietary protocol described below for 12 weeks after which weight,

nitrogen and energy gains, and carcass characteristics (% dressed weight, hepatosomatic

index) were determined. The comparative carcass analysis approach described by Cho

and Kaushik (1990) was used and provided the basis for an energy budget. In the fasting

trial, the fish were fed according to the same dietary protocol for six weeks and were then

deprived of food for two weeks. Ten fish were sampled before and after the period of

fasting to measure fasting energy (heat) losses (FHP; Blaxter 1989). In the respirometry

trial, the fish were fed according to the dietary protocol used in the feeding trial for a

minimum of four weeks before the beginning of the respirometry measurements. Each

tank of fish was allowed successively for a seven-day period (Figure 4.1) in one of two

fish respirometers similar to the one described by Cho et al. (1982).

Dietary treatments

A high fish meal diet, containing approximatively 14% digestible carbohydrate,

was used as a base for all the diets in this study. This basal diet was progressively diluted

with gelatinized potato starch (Matsunorin, Matsutani Chemical Co., Hyogo, Japan) in

the following fashion (ratio of basal diet to gelatinized potato starch) : 65:00, 65:20 and

65:35. The ingredients and chemical compositions of the diets are presented in Table 4.1

and 4.2. The diets were prepared as described in Chapter Three.

The three diets were pair-fed, block-wise, on the basis of the feed intake of the

fish fed the diet containing the highest level of starch (65:35). Those fish were fed to

near-satiety (the closest equivalent of ab libitum feeding) three times daily and their

intake represented 100%. The other diets were then fed at different levels in a manner

allowing all the tanks of fish in the same block to receive the same amount of the basal

diet. Therefore, the undiluted basal diet (65:00) was fed at 65% of the dietary intake of

the 65:35 diet and served as control. The intermediate diet (65:20) was fed at 85% of the

dietary intake of the 65:35 diet. The basal diet was also fed at the same level as the

65:35 diet to groups of fish that served as a satiation control (100:00).

Figure 4.1 Schematic representation of the experimental schedule.

Legend is as follows: w = weighing of the fish, 1a = sub-block, first set of two tanks of

block 1 in respirometers (tank #1 of block #1 in respirometer 1, tank #2 of block #1 in

respirometer 2), 1b = second set of two tanks of block 1 in respirometer (tank #3 of block

#1 in respirometer 1, tank #4 of block #1 in respirometer 2), 2a = first set of two tanks of

block 2 in respirometers (tank #1 of block #2 in respirometer 1, tank #2 of block #2 in

respirometer 2), etc.

Table 4.1 Formulation of the experimental diets.

Diets1

Ingredients (g) 65:00 65:20 65:35 100:00

Fish meal, herring, 68% CP 45.5 45.5 45.5 70.0

Fish oil 6.5 6.5 6.5 10.0

Celite AW5212 1.3 1.3 1.3 2.0

Vitamin premix3 0.65 0.65 0.65 1.0

Mineral premix4 0.65 0.65 0.65 1.0

Starch, potato, gelatinized 10.4 30.4 45.4 16.0

Total 65 g 85 g 100 g 100 g

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gelatinized starch, 65:35 = 65 parts basal diets, 35 parts gelatinized starch, 100:00 = 100 parts basal diet, 0 part gelatinized starch. 2 Celite is acid-washed diatomaceous silica3 provides per kg of basal diet: Retinyl acetate, 3750 IU; Cholecalciferol, 3600 IU; All-rac--tocopheryl acetate, 45 IU; Menadione sodium bisulfite, 15 mg; 1- Ascorbic acid, 600 mg; Cyanocobalamine, 0.0225 mg; d-biotin, 0.225 mg; Choline chloride, 1500 mg; Folic acid, 7.5 mg; myo-Inositol, 225 mg; Niacin, 225 mg; d-Calcium pantothenate, 45 mg; Pyridoxine.HCl, 15 mg; Riboflavin, 10.5 mg; Thiamin.HCl, 4.5 mg.4 Provides per kg of basal diet: Sodium chloride, 3000 mg/kg; Ferrous sulfate, 63 mg/kg; Manganese sulfate, 86 mg/kg; Zinc sulfate, 144 mg/kg; Copper sulfate, 25 mg/kg; Potassium iodide, 8 mg/kg.

Table 4.2 Chemical composition of the experimental diets.

Diets1

Composition (as is)2 65:00 65:20 65:35 100:00

Dry matter (DM), % 95.0 94.0 93.9 95.0

Crude protein (CP), % 43.4 34.5 29.7 43.4

Digestible protein (DP), % 39.5 31.5 27.0 40.0

Lipid, % 16.3 12.6 11.0 16.3

Ash, % 14.7 10.1 9.3 14.7

Nitrogen free extract (NFE), % 20.7 36.9 43.9 20.7

Gross energy (GE), MJ/kg 20.7 19.7 19.2 20.7

Digestible energy (DE), MJ/kg 19.4 17.8 16.8 19.0

DP:DE ratio, g/MJ 20.4 17.7 16.1 21.1

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gelatinized starch, 65:35 = 65 parts basal diets, 35 parts gelatinized starch, 100:00 = 100 parts basal diet, 0 part gelatinized starch.2 Dry matter, crude protein, ash according to standard AOAC (1995) methods. Lipid content determined to method of Bligh and Dyer (1959). Nitrogen-free extract obtained by difference. Gross energy by bomb calorimetry. Digestible protein = crude protein * apparent digestibility of protein (Table 4.5) and digestible energy = gross energy * apparent digestibility of gross energy (Table 4.5).

Fish and husbandry

Rainbow trout were obtained from a commercial fish farm (Spring Valley Trout

Farm, Petersburg, Ontario). One hundred fish were stocked in each of 36 fibreglass

tanks (60 L). Each tank was considered on experimental unit. The aquatic system was a

flow-through design with a flow rate of 3 L/min. Water was supplied as a mixture of

well and city water. Water temperature was maintained at 15C by injecting hot water

into the incoming waterline. Each tank was individually aerated. Photoperiod provided

was 12 h lighting and 12 h darkness using cool-white fluorescent light in a windowless

laboratory. The animals were treated in accordance with the guidelines of the Canadian

Council on Animal Care (CCAC 1984) and the University of Guelph Animal Care

Committee.

Feeding trial

The main trial was conducted over a 12-week period. The digestibility-

comparative carcass analysis protocol described by Cho and Kaushik (1990) was used for

this study. Fish were weighed every four weeks. Mortality, feed intake, and other

observations were recorded daily. Daily growth coefficient (DGC%) was calculated

according to Iwama and Tauz (1981) as follows:

At the end of the experiment, five fish per tank were randomly collected and

euthanized with a cephalic blow. The fish were rapidly blotted dry and weighed

individually. The liver was dissected and weighed. The viscera (heart, spleen,

gastrointestinal tract, swimbladder) were removed and the weight of the dressed carcass

of the fish was recorded. The hepatosomatic index (HSI%, liver weight to livebody

weight ratio) and the dressed carcass percentage (gutted carcass to livebody weight) were

calculated.

Another five fish from each tank were randomly sampled and placed in

previously weighed grinding jars. The jars were lightly capped and autoclaved at 110 C

for 25 min. After cooling, a drop of liquid antioxidant (Ethoxyquin, Monsanto Canada,

Mississauga, Ontario) was added to each jar. The autoclaved fish were then ground into

a homogeneous slurry with a blender until homogeneous. The ground samples were

transferred to shallow dishes, frozen and subsequently lyophilized. These samples were

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then reground and stored at -20C to await analysis.

Fasting trial

The fasting trial had the same experimental design as the main feeding trial. The

fish were fed according to the same dietary protocol as the fish in the main feeding trial

for 6 weeks and were then deprived from food for 14d. Five fish were randomly

sampled from each tank before (d=0) after (d=14) the period of food deprivation to

measure fasting heat losses (FHP). Fish carcass samples were processed as described

earlier. Protein and energy of the carcass samples were determined using the methods

described earlier. FHP values were corrected for losses of energy as nitrogenous

compounds (ammonia and urea) of the fish (Emmans 1994). This correction factor was

small, equivalent to about 1% of the FHP.

Respirometry trial

Each group of fish was allowed one week in one of two respirometers, as

described by Cho et al. (1975), which allow the measurement of oxygen consumption by

resting fish in standard experimental tanks. The tank was covered to prevent any

exchange of oxygen between the water surface and the atmosphere. The incoming water

was aerated in a mixing and reservoir tank to maintain a high dissolved oxygen level and

was then pumped into the fish tank through a filter and flowmeter. Dissolved oxygen

was measured using YSI dissolved oxygen meters (Model 58, Yellow Springs Instrument

Co, Yellow Springs, Ohio) and YSI dissolved oxygen probes (Model 5739, Yellow

Springs Instruments, Yellow Springs, Ohio). The output signals from the dissolved

oxygen meters, and electronic thermometers were fed into a laboratory data logger

(DataTaker Model DT-100, Data Electronics, Box Hill, Victoria, Australia) linked to a

computer (Compaq Portable II, Compaq Corporation Ltd, Houston, Texas) which was

used to display and store the data. Outputs from the instrument were read by the data

logger every 5 seconds. Hourly averages of 5 sec readings were used for the

calculations. The difference in the dissolved oxygen concentration between the in-flow

and out-flow was multiplied by the water flow rate to calculate the rate of oxygen

consumption by the fish.

The operating parameters for the respirometry trial are shown in Table 4.3. The

7-day schedule allowed one day of acclimation (fed), 3 days of observations on the fish

under fed conditions, followed by three days of fasting to estimate basal metabolism for

each of the determinations.

Oxygen consumption associated with basal metabolism was adjusted for

metabolic weight using a scaling coefficient of 0.824 as recommended by Cho and

Kaushik (1990). Oxygen consumption was used to estimate fasting heat production

(FHP) using an oxycalorific coefficient of 13.6 kJ/g oxygen consumed (Cho et al. 1982).

Oxygen consumption associated with heat increment of feeding was calculated as

the average oxygen consumption of the fed fish minus the oxygen consumption of these

fish on the 3rd day of food deprivation (oxygen consumption associated with basal

metabolism). The oxygen consumption associated with heat increment of feeding was

expressed as mg O2 per kJ basal diet consumed.

Table 4.3 Experimental conditions for respirometry.

Parameters Conditions

Fish Rainbow trout, acclimatized to diet for

minimum of 4 weeks prior to passage in

respirometer

Activity level Resting

Initial body weight 20 - 60 g/fish

Total biomass 1.8 - 4.0 kg

Water volume 30 L (50 x 30 x 20 cm)

Water flow rate 4 - 6.5 L/min

Water temperature 15C

Dissolved oxygen in incoming water 8.0 - 10 mg/L

Lighting 12 h light : 12 h dark

Feeding 2 times daily

Data collection 1 reading / min

1 reading / h (average of 5sec readings)

Experimental period 1 d acclimation (fed)

3 d feeding

3 d fasting

Apparent Digestibility

Apparent digestibility coefficients of diets were measured using the method of

Cho et al. (1982) described in Chapter Three. Acid-washed diatomaceous silica (Celite

AW 521, Celite Corp., Lompoc, CA), a source of acid-insoluble ash (AIA), was added to

the diets to serve as a digestion indicator (Atkinson et al. 1984). A first series of fecal

samples was centrifuged after collection from the feces settling column. The highly

digestible nature of the basal diet (high fish meal, little plant products) led to the

collection of only a quantity of excreted feces with poor stability since they contained

relatively little organic matter and had a high ash content. The feces appeared less stable

in water than feces samples obtained with the reference diet of Cho et al. (1982). The

fecal particles were very fragile and were sensitive even to the limited handling prior to

centrifugation. Centrifugation at 3000 xG for 20 min at 4C led to the physical

separation of the undigested starch from the rest of the feces. The loss of a significant

amount of what appeared to be starch "dust" during freeze-drying was also noticed

(during building and release of the vacuum in the freeze-drier). Apparent digestibility

coefficients of dry matter, energy and protein from these samples were very high

denoting significant loss. The results from this set of samples were, therefore, not used

for this study and are not presented.

A second set of feces samples was, therefore, collected very carefully from the

feces collector and the "intact" feces suspended in a large quantity of water were frozen

immediately to -10C and subsequently freeze-dried as is. The freeze-dried fecal samples

from two collection periods had to be pooled because of the very small amount of feces

collected from the fish fed the 65:00 diet. Since only the results from one pooled sample

were available, no statistical analysis on the results from the digestibility trial could be

performed.

Chemical analyses

Samples of diet, feces and carcass were analyzed via Weende proximate analysis.

Diets, ingredients, feces and dry carcass were analyzed for dry matter (DM), crude

protein (%N x 6.25) by the Kjeldahl method using a Kjeltech 1030 autoanalyzer

(Tecator, Hoganas, Sweden), ash according to AOAC (1995), and total lipid according to

the method of Bligh and Dyer (1959). Gross energy content of carcass samples was

measured using a Parr 1271 automated bomb calorimeter (Parr Instruments, Moline,

Illinois). Acid insoluble ash (AIA) was analyzed according to Atkinson et al. (1984b).

Energy Budget

The analysis of the results obtained from the comparative carcass analysis were

integrated using the energy budget presented in Figure 2.1. HjE (activity) was assumed

to be minimal since visual observation of the fish through a video camera showed that

they rested for most of the time on the bottom of the tank. HjE was also assumed to be

similar for fed and unfed fish. The HjE was, therefore, confounded in the HeE estimate.

The information from the three trials was integrated to build, step by step, a detailed

energy budget. Calculations for some of the components of the energy budget are

described in the legends of certain tables. Estimation of HeE for the energy budget of

the fish in the feeding trial (Equation 4.3) was estimated from FHP calculated in the

fasting and respirometry trials. Metabolic body weight (MBW) of the fish was estimated

for each day of the experimental period (total of 84 days) by recreating their growth

curve using the DGC growth model presented earlier in Equation 4.1. For each of the 84

days of the experiment, MBW was multiplied by the estimate of HeE obtained from the

fasting and respirometry trials as follows:

Digestible, metabolizable and net energy were calculated as described by Cho and

Kaushik (1990) and the NRC (1981).

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Statistical Analysis

Dependent variables (and composite variables with a strong dependent

component, e.g. feed efficiency) were analyzed using the general linear model (GLM)

with the SAS/STAT software (SAS 1988) using a complete block design. Means were

compared using Tukey's honestly significant difference test with an = 0.05 (Steel and

Torrie 1980).

4.3 RESULTS4.3 RESULTS

Feeding trial

The feed and nutrient (protein, lipid, and nitrogen-free extract) intakes of the fish

in the feeding trial are given in Table 4.4. The apparent digestibility coefficients (ADC)

of the diets are presented in Table 4.5. ADC of protein was high which is an indication

that the fish meal used was of good quality. ADC of energy of diet 65:00 and 100:00

were high, but digestibility of energy appeared to decrease slightly with an increase in

the level of dietary starch.

Table 4.4 Calculated nutrient intakes per fish over a 12-week experimental period.

Diet1

Intake2 65:00 65:20 65:35 100:00

Feed, g 39.7 50.9 58.7 59.7

Dry matter, g 37.7 47.8 55.1 56.7

Crude protein, g 17.2 17.6 17.4 25.9

Digestible protein, g 15.7 16.1 15.8 23.9

Lipid, g 6.5 6.5 6.5 9.7

Nitrogen-free extract, g 8.2 18.8 25.8 12.4

Gross energy, kJ 823 1003 1125 1237

Digestible energy, kJ 770 905 983 1138

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gelatinized starch, 65:35 = 65 parts basal diets, 35 parts gelatinized starch, 100:00 = 100 parts basal diet, 0 part gelatinized starch.2 Intake = Feed intake per fish per 84 d * composition. Dry matter, crude protein, ash according to standard AOAC (1995) methods. Lipid content determined to method of Bligh and Dyer (1959). Nitrogen-free extract obtained by difference. Gross energy by bomb calorimetry. Digestible protein = crude protein * apparent digestibility of protein (Table 4.5) and digestible energy = gross energy * apparent digestibility of gross energy (Table 4.5).

Table 4.5 Apparent digestibility coefficients (ADC) of experimental diets (based on

pooled samples from 2 weeks collection, n=1).

Diets1

ADC 65:00 65:20 65:35 100:00

Dry matter,% 84.6 87.5 87.3 86.3

Crude protein, % 91.1 91.2 90.9 92.1

Energy, % 93.5 90.2 87.4 92.0

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gelatinized starch, 65:35 = 65 parts basal diets, 35 parts gelatinized starch, 100:00 = 100 parts basal diet, 0 part gelatinized starch.

Figure 4.2 shows the growth curve of fish fed the experimental diets. Table 4.6

summarizes the fish growth performance, carcass characteristics and apparent nitrogen

and energy utilization. Fish fed the 100:00 diet had a significantly higher weight gain

and growth rate (DGC%) than the fish fed the other diets. The weight gain of fish fed

the 65:20 diet was slightly, yet significantly, greater than those of fish fed the 65:00 and

65:35 diets. However, the dressed carcass weight of fish fed the 65:20 diet was not

significantly different from that of fish fed the 65:00 diet. The greater live weight of fish

fed the 65:20 diet compared to the fish fed the 65:00 diet was due to the greater size of

their liver. The relative size of the liver of the fish fed the 65:35 diet was significantly

greater than that of the fish fed the 65:00, 65:20 and 100:00 diets. The fish fed the 65:35

diet had a live weight not different from the fish fed the 65:00 diet, but had a

significantly lower dressed carcass weight. Feed efficiency was not different for fish fed

the 65:00 and 100:00 diets. Feed efficiency decreased significantly with an increase in

gelatinized starch intake.

Figure 4.2 Growth curve of the fish fed the experimental diets for 12 weeks.

Diet designation are as follows: 65:00 = 65 parts basal diet, 0 part gelatinized starch,

65:20 = 65 parts basal diet, 20 parts gelatinized starch, 65:35 = 65 parts basal diets, 35

parts gelatinized starch, 100:00 = 100 parts basal diet, 0 part gelatinized starch

Table 4.6 Performance and carcass characteristics of the fish fed the experimental diets over a 12-week period1.

Diets2

Parameters 65:00 65:20 65:35 100:00 SEM HSD3

Feed, g/fish 39.7 50.9 58.7 59.7 0.4 N/A4

Gain, g/fish 52.7c 55.8b 51.6c 79.0a 0.3 1.6

FE, gain:feed 1.33a 1.10b 0.90c 1.32a 0.01 0.05

DGC, % 2.29c 2.37b 2.26c 2.89a 0.01 0.04

HSI, % 1.56c 4.46b 5.68a 1.55c 0.2 1.1

Dressed carcass, % 87.4a 84.3b 81.6c 86.8a 0.5 2.3

Dressed carcass, g/fish 53.1b 53.9b 48.7c 75.6a 0.4 1.8

Retained N, g/fish 1.26b 1.35b 1.21b 1.98a 0.04 0.07

Retained N, % of intake 45.7a 49.5a 44.8a 46.2a 1.2 5.8

RE, % DE 48.9b 45.3c 38.0d 55.9a 0.7 3.2

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gel. starch, 65:35 = 65 parts basal diets, 35 parts gel. starch, 100:00 = 100 parts basal diet, 0 part gel. starch.2 Initial body weight = 8.1 g/fish3 Tukey honestly significant difference. Means in the same row sharing the same letter are not statistically different.4 Not applicable, independant variable

Carcass composition data is presented in Table 4.7. Fish fed diet 100:00

contained a significantly lower percentage of moisture and a higher percentage of lipid

than the fish fed the other diets. Fish fed the 65:35 had a significantly lower protein

content than the fish fed the 65:00, 65:20 and 100:00 diets. The fish fed the 65:00 and

100:00 diets appared to have no or very little NFE (calculated by difference). NFE,

however, appear to increase with increasing digestible starch intake.

The energy budget calculated for the fish of the feeding trial is presented in Table

4.8. The fecal energy losses (FE) increased with increasing gelatinized starch intake.

Non-fecal losses (UE+ZE), derived solely from calculations based on N excretion, were

not significantly different among diets 65:00, 65:20 and 65:35. Fish fed the 100:00 diet

had significantly greater estimated non-fecal energy losses. Fish fed the 100:00 diet had

much greater RE than fish fed all other diets. The fish fed the 65:20 diet also had

significantly higher RE than the fish fed the 65:00 and 65:35 diets, although this did not

extend to the dressed carcass.

Table 4.7 Whole body carcass composition of the fish fed the experimental diets.

Diets1

Composition2 65:00 65:20 65:35 100:00 SEM3 HSD4

Moisture, % 73.9a 73.3a 73.5a 71.9b 0.2 0.9

Crude protein, % 14.9a 15.0a 14.4b 14.9a 0.1 0.5

Lipid, % 9.3b 9.4b 9.3b 11.0a 0.1 0.4

Ash, % 2.0a 2.0a 2.0a 2.2a 0.04 0.2

Nitrogen-free extract, % 0.0b 0.3ab 0.8a 0.1b 0.1 0.5

Gross energy, kJ/g 7.3b 7.4b 7.4b 8.1a 0.1 0.3

1 Diet designation: 65:00 = 65 parts basal diet, 0 parts gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gel. starch, 65:35 = 65 parts basal diets, 35 parts gel. starch, 100:00 = 100 parts basal diet, 0 part gel. starch.2 Dry matter, crude protein, ash according to standard AOAC (1995) methods. Lipid content determined to method of Bligh and Dyer (1959). Nitrogen-free extract obtained by difference. Gross energy by bomb calorimetry.3 Tukey honestly significant difference. Means in the same row sharing the same letter are not statistically different.4 Not applicable, independant variable

Table 4.8 Energy budget of the fish fed the experimental diets for a 12-week period in the main feeding trial.

Components Diets1

kJ/ fish per 84d 65:00 65:20 65:35 100:00 SEM HSD

IE 823 1003 1125 1237 N/A N/A

FE 54c 94b 142a 99b 0.4 2

UE+ZE2 37b 35b 37b 54a 1 6

RE 376c 410b 373c 635a 5 22

Balance3 355c 460b 573a 449b 10 47

1 Diet designation: 65:00 = 65 parts basal diet, 0 parts gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gel. starch, 65:35 = 65 parts basal diets, 35 parts gel. starch, 100:00 = 100 parts basal diet, 0 part gel. starch.

2 based on calculated nitrogen losses3 Obtained by difference (IE-FE-(UE+ZE)-RE), therefore, includes HE, as well as other unmeasured losses, and the errors from various other measurements.

Fasting trial

Weight gain of the fish of the fasting trial for the 6 weeks prior to the fasting

period (Table 4.9) followed the same pattern as the weight gain of the fish in the feeding

trial. Fish fed the 100:00 diet had significantly higher weight gain than the fish fed the

other diets. The weight gain of the fish fed the 65:20 diet was significantly greater than

that of the fish fed the 65:00 and 65:35 diets. Weight, nitrogen and energy losses during

the 14d food deprivation period of the fish fed the 65:00, 65:20 and 65:35 were not

statistically different. Fish fed the 100:00 diet had a significantly greater weight loss

than the fish fed the 65:00, but nitrogen and energy losses that were not significantly

different from those of the fish fed the 65:00, 65:20 and 65:35 diets. FHP calculated

from the carcass energy loss of the fish in the fasting trial was not statistically different

among diets. FHP values presented in Table 4.9 were used as estimates of HeE for the

energy budget of the fish from the feeding trial in Table 4.11.

Table 4.9 Weight, nitrogen, and energy losses and calculated fasting heat losses of rainbow trout during fasting trial.

Diets1

Parameters 65:00 65:20 65:35 100:00 SEM HSD

Weight gain2, g/fish (6 wks feeding) 18.3c 19.8b 18.8c 27.8a 0.2 0.9

Weight loss, g/fish*day-1 0.21b 0.27ab 0.28ab 0.31a 0.02 0.08

N loss, mg/fish*day-1 2.7a 3.6a 3.1a 4.5a 0.8 3.8

Energy loss3, kJ/fish*day-1 1.95a 2.69a 1.88a 2.46a 0.34 1.67

FHP, kJ/kg0.824*day-1 39.5a 52.4a 37.7a 38.3a 6.1 30

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gel. starch, 65:35 = 65 parts basal diets, 35 parts gel. starch, 100:00 = 100 parts basal diet, 0 part gel. starch.

2 Initial live bodyweight = 8.1 g/fish3 corrected for nitrogenous energy losses (24.9 kJ/g N lost)

Respirometry trial

FHP values calculated from oxygen consumption of the fish on the third day of

fasting in the respirometry trial (Table 4.10) were not significantly different among diets.

The FHP values from the respirometry trial were slightly lower than FHP obtained from

the fasting trial. Oxygen consumption values associated with heat increment of feeding

(HiE), which was calculated after subtraction of the oxygen consumption of fasted fish

were not significantly different among diets.

Figure 4.3 presents an example of the oxygen consumption of the fish fed one of

the experimental diets (65:20 diet, data from one day only) and the oxygen consumption

of the same fish on the third day of fasting.

Table 4.10 Oxygen consumption and calculated heat production of fish measured in the respirometry trial on the 3rd day of food

deprivation (FHP) or attributed to heat increment of feeding (HiE).

Diets1

Components 65:00 65:20 65:35 100:00 SEM HSD2

FHP, g O2/kg0.824 per day 2.29a 2.78a 2.20a 2.86a 0.36 1.88

FHP, kJ/kg0.824 per day3 31.2a 37.9a 30.0a 39.0a 4.9 25.7

HiE, mg O2 per kJ IE basal diet 9.33a 8.21a 10.70a 6.69a 1.4 7.35

HiE, kJ/kJ basal diet3 0.13a 0.11a 0.15a 0.09a 0.02 0.10

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gel. starch, 65:35 = 65 parts basal diets, 35 parts gel. starch, 100:00 = 100 parts basal diet, 0 part gel. starch.

2 Tukey honestly significant difference. Means in the same row sharing the same letter are not statistically different.3 Based on an oxycalorific coefficient of 13.6 kJ/O2 consumed.

Figure 4.3 Example of the measured oxygen consumption of fish (oxygen

consumption expressed per fish basis, average weight = 39 g/fish) fed the 65:20 diet on

the 2nd day of feeding and on the 3rd day of fasting.

Two estimates of HeE were calculated for the fish of the main feeding trial using

FHP values calculated either from the fasting trial or from the respirometry trial (Table

4.11). HiE was also calculated for the fish of the main feeding trial based on the estimate

of HiE obtained by respirometry. This led to deviations designated on Table 4.11 which

varied widely among diets and ranged between 8 - 27% the equivalent of IE. The

calculated heat production for the fish for the feeding trial (Table 4.11) showed a slight,

but significant, increase with increasing starch intake. The calculated increases in heat

production represented only about 5% of the gross energy intake (IE) provided as starch.

Energy values of the 65:00 and 100:00 diets were very similar (Table 4.12). The

65:20 and 65:35 diets showed DE and ME values relatively close to diet 65:00 and

100:00. The NE content of 65:20 diet was relatively close to that of the 65:00. The

65:35 diet, on another hand, had much lower NE value than the 65:00 and 100:00 diets.

Table 4.11 Estimated heat losses per fish for 12-week main feeding trial.

Diets1

Component 65:00 65:20 65:35 100:00 SEM HSD2

Balance feeding trial3 355c 460b 573a 449b 10 47

HeE fasting trial4 184c 252a 175d 226b 0.6 3

HeE respirometry trial4 141c 176b 134d 220a 0.3 2

HiE respirometry trial 5 105c 79d 139a 113b 1 5

HE fasting trial6 289d 331b 314c 339a 1 7

HE respirometry trial7 246d 255c 273b 333a 2 8

Deviation I 8 66c 129b 258a 109b 9 42

Deviation II 9 110c 205b 300a 115c 9 42

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gel. starch, 65:35 = 65 parts basal diets, 35 parts gel. starch, 100:00 = 100 parts basal diet, 0 part gel. starch.

2 Tukey honestly significant difference. Means in the same row sharing the same letter are not statistically different.3 from Table 5.7, obtained by difference: (IE-FE-(UE+ZE)-RE) feeding trial4 calculated according to Equation 4.3 5 calculated from estimate of HiE obtained from respirometry trial (Table 5.9)6 HeEfasting trial+HiErespirometry7 HeErespirometry trial +HiErespirometry trial8 obtained by difference: Balance feeding trial - (HeE fasting trial + HiErespirometry trial)9 obtained by difference: Balance feeding trial - (HeE respirometry trial + HiErespirometry trial)

Table 4.12 Calculated energy values of the experimental diets.

Diets1

Energy value 65:00 65:20 65:35 100:00

Gross energy (GE), MJ/kg 20.7 19.7 19.2 20.7

Digestible energy (DE), MJ/kg 19.4 17.8 16.8 19.0

Metabolizable energy (ME), MJ/kg 18.4 17.2 16.1 18.1

Net energy (NE), MJ/kg2 14.1 13.0 9.4 14.5

Net energy (NE), MJ/kg3 13.0 11.4 8.6 14.2

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gel. starch, 65:35 = 65 parts basal diets, 35 parts gel. starch, 100:00 = 100 parts basal diet, 0 part gel. starch.

2 based on HeEfasting trial (Table 4.11)3 based on HeErespirometry trial (Table 4.11)

4.4 DISCUSSION4.4 DISCUSSION

The pair-feeding technique used in this study was a practical and very effective

method of determining the energy value of carbohydrate. Fish accepted the diet with the

highest starch very well. Growth rates (DGC) observed in this study were within the

range of growth rates routinely obtained with rainbow trout at the University of Guelph.

No statistical analysis could be performed on the result of the digestibility trial

because of the limited amount of sample collected. Nonetheless, the results appear to be

reliable, since the ADC of protein measured are in accordance with what is routinely

observed in this laboratory for good quality herring meal. The ADC of energy of the

diets are also in line with ADC of energy of diets containing gelatinized starch obtained

in this laboratory (Cho and Kaushik 1990) and elsewhere (Kim 1989).

The results from the respirometry trial suggest that an increase in digestible

carbohydrate intake did not lead to any significant increase in oxygen consumption of

fish and may indicate that the supplemental carbohydrate was not (aerobically)

catabolized by the fish. The calculated heat production of the fish of the feeding trial in

Table 4.11 showed a slight increase with increasing starch intakes, but this increase

represented less than 10% of the gross energy intake provided as supplemental starch.

Thus, the heat production of the fish, estimated from oxygen consumption, was not

greatly affected and could only account for a small portion of the ingested (supplemental)

carbohydrate energy. Since heat production was estimated based on oxygen consumption

and there is possibility of real heat losses differed than those estimated in the present

study since oxygen consumption does not allow the estimation of heat losses associated

with anaerobic catabolism (Brafield and Llewellyn 1982).

Feeding digestible starch to rainbow trout had a small but significant effect on

growth and RE in the fish fed the 65:20 diet compared to those fed the 65:00 diet but not

in the fish fed the 65:35 diet. The greater weight and energy gains of the fish fed the

65:20 diet compared to the 65:00 diet was due to increased weight of the viscera (liver

included), since the fish fed these two diets did not differ in dressed carcass weights.

Fish fed the 65:35 diet did not show any increase in growth or recovered energy and even

showed a decrease in dressed carcass weight compared to fish fed the 65:00 diet. This

difference in response of the fish to the 65:20 and 65:35 diets is difficult to explain. It is

probably not due to experimental error because similar results were obtained in the 6-

week feeding period prior to the 14d food deprivation in the fasting trial (Table 4.9) and

in another study which used the same dietary protocol (Page 1996). At an intermediate

digestible starch intake, therefore, the fish appeared to deal with the influx of

carbohydrate by storing part of it in their liver (shown by a significant increase in the size

of the liver) and this was without effect on growth of the fish in terms of carcass yield.

At high starch intake (65:35), the liver size of the fish increased even further, and this

may have affected carcass yield. This higher weight of the liver may have required a

significant channeling of amino acids toward hepatic protein synthesis. The amount of

amino acids necessary to sustain growth of the liver may have reached a critical level for

the fish fed the 65:35 diet and may, consequently, have affected synthesis of carcass

protein by these fish. This is only, however, only hypothetical since the composition of

the liver was not determined in the present study. Beside a slight effect on carcass yield

in fish fed the 65:35 diet, increasing digestible carbohydrate intake did not appear to

create any ill-effects, at least for the short duration of the feeding trial.

The lack of significant difference among diets 65:00, 65:20 and 65:35 in terms of

RN at similar digestible protein intakes indicates that the supplemental starch had no

sparing effect on dietary protein. The results appear to be in contradiction with the

results of Kim (1989) and Brauge (1994), who concluded that, even at high intakes,

carbohydrate contributed positively to growth and recovered energy and had a protein-

sparing effect in rainbow trout.

The various components of the energy budget quantified accounted for

approximately 90% of the IE of the fish fed the 65:00 and 100:00 diets, leaving

deviations (unquantified energy losses) of about 10% (Table 4.11). The deviations for

the fish fed the 65:00 and 100:00 can be attributed to inevitable energy losses, such feed

wastage, energy losses as in tegument and mucus that were not quantified and various

errors associated with the several components of the energy budget. The deviations,

nonetheless, were increased significantly by feeding the supplemental starch, indicating

that other significant sources of energy losses remain still unindentified and unquantified.

The conclusion of Kim (1989) and Brauge (1994) that digestible carbohydrate, at

high levels of intake, are an efficient source of energy appear in contradiction with the

results from the present study. The similarity between the growth performances and feed

efficiencies of fish fed diets of similar composition in studies presented here and the

studies of Kim (1989), and Brauge (1994) suggests that the conclusion of Kim (1989)

and Brauge (1994) differs from the conclusion reached in the present study because of

differences in the interpretation of the results or in the basis for comparison (i.e. the

control diets). Kim (1989), for example, used a control diet containing 38% raw (poorly

digestible) starch and low level of lipids (10%). This control diet supported suboptimal

growth performance and low nitrogen retention efficiency. A high digestible

carbohydrate diet supported slightly better performance than the control diet. The high

digestible carbohydrate diet had a composition similar to the 65:35 diet used in the

present study and both diets supported similar growth rates and feed efficiencies. The

conclusion of Kim (1989) that digestible carbohydrate, at high levels of intake, is as

efficient a source of energy and that dietary carbohydrate can spare a significant amount

of protein may only be valid at low lipid intake and at suboptimal growth performances

levels (where important catabolism of protein is occurring to meet energy requirement).

The results from the present study do not suggest any glucostatic control of feed

intake (Brafield and Llewellyn 1982). Hilton et al. (1987) suggested that such a

mechanism could be present in fish based on the result of their study with rainbow trout.

The present study strongly suggests, however, that the suboptimal growth performance of

fish fed high levels of carbohydrate is due to the low net energy (NE) content of such

diets. In the present study, dilution of the diet with starch resulted in a decrease in the

NE of the diet. Cho and Woodward (1989) concluded that diets with digestible energy

contents of less than 15 MJ/kg could not support optimal growth in salmonids because,

below this level, the fish could not consume enough diet. A DE level of 15 MJ/kg

translates into 12-13 MJ/kg NE for a good quality salmonid diet fed to fish under normal

culture conditions (Cho and Kaushik 1990). The basal diet (65:00 and 100:00) had a NE

value of about 14 MJ/kg, while the 65:35 diet had a NE value of only about 9 MJ/kg

(Table 4.12). The latter level is probably too low to allow the fish to compensate by

increasing intake and the low nutrient density and energy density probably prevented the

consumption of enough nutrients and NE to support higher growth. The NE of the 65:20

diet was between 11.4 and 13 MJ/kg which are to close to the estimated minimum NE

required based on the conclusions of Cho and Woodward (1989). The feed intake of the

fish fed the 65:20 diet was restricted in the present study, it is not known if the 65:20 diet

can support optimal growth performance. The estimate of NE of 65:20 may only be true

for the particular conditions used in this present study, since our results show that there is

no fixed NE value for carbohydrate (i.e. carbohydrate contributes little NE in excess of a

certain intake level). The NE of the 65:20 diet could have been lower if this diet had

been fed to near-satiety.

The ability of the fish to compensate for dilution of the NE of the diet will

probably depend on several factors. Whether the diet is diluted with a non-digestible or

digestible component (both of low NE value) is one factor influencing the ability of the

fish to compensate, since digestible components could clear the gastro-intestinal tract

more rapidly.

The fasting heat losses (FHP) measured by carcass analysis following 14d of

fasting varied between 37.7 and 52.4 kJ/kg0.824 per day (mean = 42.0 kJ/kg0.824 per day).

FHP, calculated from oxygen consumption of the fish on the 3rd day of food deprivation,

varied between 30.0 - 39.0 kJ/kg0.824 per day (mean= 34.5 kJ/kg0.824 per day). The FHP

obtained from the fasting trial (14d fasting) as well as those obtained from the

respirometry trial were, therefore, similar to the results of Cho and Kaushik (1990), who

estimated FHP to be around 40 kJ/kg0.824, based on oxygen consumption of rainbow trout

of various size at 15C. The results of the present study are also comparable to the

estimates of Kaushik and Gomes (1988) who measured values between 32.9 - 43.0

kJ/kg0.824 per day at 18C with 150 g rainbow trout. These FHP values are, however,

considerably lower than the values obtained by Smith et al. (1978a) using direct

calorimetry. Direct calorimetry probably does not allow accurate quantification of heat

losses in fish because of practical obstacles, such as the small amount of heat release by

the animal. The latter authors estimated FHP at 15C to be 204 kJ/kg0.75 body weight for

1-57 g rainbow trout. The very good agreement between the results of Kaushik and

Gomes (1988), Cho and Kaushik (1990) and the present study suggest that FHP of

rainbow trout (and probably also other salmonids) can roughly be estimated at 40

kJ/kg0.824 per day at 15C. This, at least, holds true for rainbow trout with BW between 1

and 150 g. There would be interest in verifying if this estimate is applicable for much

larger fish.

Chapter FIVE. URINARY EXCRETION OF GLUCOSE BY RAINBOW TROUT

AT INCREASING DIGESTIBLE STARCH INTAKESChapter FIVE. URINARY

EXCRETION OF GLUCOSE BY RAINBOW TROUT AT INCREASING

DIGESTIBLE STARCH INTAKES

5.1 INTRODUCTION.1 INTRODUCTION

Analysis of data from feeding trials shows that at high carbohydrate intake a large

portion of the energy absorbed by the fish is unaccounted for when comparing intake of

energy and recovered energy in the carcass, fecal energy losses, non-fecal nitrogenous

energy losses and heat production estimated from oxygen consumption (Chapter Four).

This discrepency in the energy budget of these fish could be due to excretion of glucose

in the urine as suggested by Furuichi (1988) and Shiau and Suen (1992).

The excretion of a portion of the dietary glucose or of its metabolites in the urine

would mean that diets containing high levels of dietary carbohydrate would have lower

ME values than calculated only on the basis of nitrogenous (ammonia and urea) energy

excretion. Quantification of urinary excretion of glucose by the fish is necessary to

obtain a clearer picture of the energy partitioning of carbohydrate by trout.

5.2 MATERIALS AND METHODS.2 MATERIALS AND METHODS

The excretion of urinary glucose was measured in rainbow trout using the

method of Curtis and Wood (1991). This method involves the determination of the urine

flow rate of non-catheterized fish using a glomerular filtration marker, 1,2 3H

polyethylene glycol 4000 (1,2 3H PEG-4000), and spot sampling of urine to measure

concentration of substances of interest.

Initially, attempts were made to decrease the volume of water of the tank from 50

L to 20 L and use one fish per tank to obtain estimates of UFR and urinary glucose

excretion rate for individual fish. Under these conditions, however, fish were highly

excitable and refused to feed or had very low feed intake. The following experimental

design was, therefore, used. Forty-eight rainbow trout (live body weight = 269 g/fish)

were randomly divided in twelve groups of four fish and were stocked in twelve

rectangular fibreglass tanks (50 L) supplied with a mixture of well water and city water

at a rate of about 3 L/min. Each tank was individually aerated, and water temperature

was controlled thermostatically at 15C. Photoperiod was maintained on a 12 h light : 12

h dark schedule. The animals were treated in accordance with the guidelines of the

Canadian Council on Animal Care (CCAC 1984) and the University of Guelph Animal

Care Committee. The four dietary treatments were each allocated to three tanks using a

complete block design. The four diets were fed according to the dietary protocol

described in Section 4.2 for an acclimation period which lasted 8 days.

After the acclimation period, the fish were then anaesthetized with tricaine

methanesulfonate (MS222) and 0.6 ml of Cortland saline containing 17 Ci of 1,2 3H

polyethyelene glycol (NET-405, Mandel Scientific Co. Ltd, Guelph, Ontario) was

injected into the caudal vein of each fish using a 26 gauge needle. This dose was

identical to that used by Curtis and Wood (1991). After the injection, the fish were then

returned to their original tanks (50 L each) and were pair-fed as previously for the

following 48-hour period.

Forty-eight hours after injection of the glomerular filtration marker, the water

flow was interrupted and each tank was stoppered with a rubber cork. Vigorous aeration

was, however, maintained in each tank. Water samples (5 ml) were taken at 0, 1, 2, 3

and 4 hour after interruption of water flow to determine the excretion of tritiated PEG-

4000 in the water and permit calculation of the urine flow rates. The water samples were

diluted with 15 ml liquid scintillation fluid (Scintiverse II, Fisher Scientific, Fairlawn,

NJ).

After the 4 h water sampling period, the fish were anaesthetized with MS 222.

Urine was spot-sampled from the urinary bladder using a polyethylene catheter (PE-50,

Clay Adams, Parsippany, NJ) attached to a 1cc syringe with a blunted 23 gauge needle.

A blood sample (500 L) was then taken from the caudal vein of each fish. The blood

was transferred to a plastic microcentrifuge tube containing 100 L heparinized saline,

mixed and immediately centrifuged in a micro-centrifuge (Model 235C, Fisher

Scientific, Fairlawn, NJ) for 10 min at 13,600xg. Urine and plasma samples (25 L)

from each fish were diluted in 15 ml of liquid scintillation fluid. Urine and plasma

samples were rapidly frozen in liquid N2 and subsequently kept at -80C to await

analysis. They were subsequently analyzed for glucose using an enzymatic

determination kit using hexokinase (Sigma, St-Louis, Mo). 3H in water, plasma and

urine samples was counted (up to 2) using a liquid scintillation counter (LS-3801,

Beckmann Instrument, Fullerton, Ca) after 12h of dark acclimation. Approximately 60%

counting efficiency was achieved.

Urine flow rate (UFR) and glomerular filtration rate (GFR) were calculated for

each group of four fish based on the concentration of 1,2 3H PEG-4000 of the urine and

on the accumulation in the tank of 1,2 3H PEG-4000 attributable to urinary excretion.

Approximately 80% of total excretion of 1,2 3H PEG-4000 is attributable to urinary

excretion according to Curtis and Wood (1991) with the 20% remaining assumed to be

attributable to branchial excretion (C.M. Wood, personal communication). The

calculations were as follows:

Install Equation Editor and double-click here to view equation.

Statistical analyses

Before analysis of variance, plasma and urinary glucose concentrations, as well as

the estimated urinary glucose excretion rates of individual fish, were subjected to

logarithmic transformation in order to improve homogeneity of variance (Lison 1968).

These log transformed variables were averaged by tank (experimental unit) and analysis

of variance was performed using the general linear model (GLM) using the SAS/STAT

software (SAS 1988) using a complete block design. Three pre-planned comparisons

(65:00 vs 65:20, 65:00 vs 65:35, 65:20 vs 65:35) were performed using orthogonal

contrasts with a level = 0.05 (Steel and Torrie 1980).

Determination of a renal threshold for glucose excretion

Urine glucose concentration (Figure 5.1) and estimated urinary glucose excretion

(Figure 5.2) were plotted against plasma glucose concentrations. Renal threshold for

glucose excretion was estimated using several approaches: 1) minimum plasma glucose

observed to produce an urinary glucose concentration superior to 0.3 mM (minimum

resolution level of the glucose assay), 2) maximum plasma glucose observed to product

Install Equation Editor and double-click here to view equation.

an urinary glucose concentration lower than 0.3 mM and 3) linear regression of urinary

glucose concentration or estimated urinary glucose excretion against plasma glucose

concentration. The latter approach was based on the theoritical relationship between

plasma glucose and urinary glucose excretion (Figure 2.3) where the urinary glucose

excretion rate increases linearly when the glucose reabsorption system is completely

saturated (Khoushanpour 1976). Linear regression analysis was performed using the

linear regression (REG) procedure of the SAS/STAT software (SAS 1988). Urinary

glucose concentration or excretion values corresponding to progressively lower values of

plasma glucose were included in the analysis in a stepwise fashion in a procedure aiming

at the minimization of the mean square error (MSE). This resulted in the incorporation

of all the data points in the regression analysis. All the data points apparently belonged

to the linear regression model. Renal threshold for glucose excretion was determined at

the plasma concentration resulting in urinary glucose concentration equal to 0 mM or

urinary glucose excretion equal to 0 mmol/kg per h. A quadratic model was also used,

but the nature of the response was almost entirely linear in nature (results not shown).

5.3 RESULTS AND DISCUSSION.3 RESULTS AND DISCUSSION

The measured UFR varied between 4.9 and 7.1 mL/kg*h-1 and the GFR ranged

between 12.7 and 17.4 mL/kg*h-1 (Table 5.1). No significant differences were observed

for the UFR and GFR of the fish fed the four experimental diets. These UFR and GFR

values are much greater than those reported by Curtis and Wood (1991), who measured

UFR of only 2.4 ml/kg*h-1 (200 - 425g fish, 15C) and GFR of 4.5 ml/kg*h-1.

The difference between the UFR and GFR measured in the present study and

those measured by Curtis and Wood (1991) may be explained in part by the fact that the

latter study used fish that were unfed for 7 days prior to the measurement. A significant

reduction in the urine production has been observed in fish during a fasting period (Hunn

1982). It may be hypothesized that fish with a higher metabolic rate produce more urine.

Based on estimates of Cho (1992), the metabolic rate of the fish from the present study

was expected to be 150-200% of that of the fish used by Curtis and Wood (1991).

Changes in surface area and blood perfusion (to increase oxygen uptake) may be major

factors influencing the flux of water through the gills (Hunn 1982, Isaia 1984). Urine

production increases with temperature with a Q10 of about 2, which is relatively

consistent with Q10 estimates based on oxygen consumption (Hunn 1982). Wood and

Randall (1973) showed that UFR rose during exercice which is in agreement with the

hypothesis that an increase in oxygen consumption increases urine production.

Plasma glucose concentrations increased significantly at high digestible

carbohydrate intake since the fish fed the 65:35 diet having a significantly higher plasma

glucose level than fish fed the 65:00 diet. (Table 5.2). Urinary glucose concentrations

varied greatly from close to zero up to concentrations several times greater than that of

plasma and did not differ significantly between diets. The fish fed the 65:35 diet,

nonetheless, tended to have greater urine glucose levels than the fish fed the 65:00

(P<0.06). The glucose excretion rate of the fish fed the 65:35 was significantly greater

than that of fish fed the 65:00 and 65:20 diets (Table 5.2).

Table 5.1. Urine flow rate (UFR) of fish fed the various experimental diets.

Diets1

65:00 65:20 65:35 100:00 SEM Comparisons2

UFR, ml/kg per h 5.5 4.9 7.1 6.6 1.0 No significant differences

GFR, ml/kg per h 17.4 12.7 13.0 13.8 1.6 No significant differences

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gelatinized starch, 65:35 = 65 parts basal diets, 35 parts gelatinized starch, 100:00 = 100 parts basal diet, 0 part gelatinized starch.

2 Comparison of means using orthogonal contrasts (65:00 vs 65:20, 65:00 vs 65:35, 65:20 vs 65:35) with =0.05.

Table 5.2 Plasma and urine glucose concentrations and urinary glucose excretion of the fish fed the experimental diets.

Diets1

Glucose Concentration 65:00 65:20 65:35 100:00 SEM Comparisons2

Plasma

Mean3, mM 5.4 9.5 20.7 10.9 4.8 65:00 vs 65:35 P<0.0565:00 vs 65:20 N.S.65:20 vs 65:35 N.S.

Range4, mM 2.2 - 7.1 4.3 - 18.6 7.0 - 53.2 3.5 - 36.7

Urine

Mean3, mM 3.2 11.2 44.7 15.3 14.2 No significant differences

Range4, mM 0.2 - 12.9 0.2 - 107.8 0.2 - 160.1 0.1 - 99.9

Glucose excretion

Mean3, mmol/kg*h-1 0.02 0.04 0.31 0.10 0.08 65:00 vs 65:20 P<0.0565:00 vs 65:35 P<0.0565:20 vs 65:35 N.S.

Range4, mmol/L 0 - 0.06 0 - 0.31 0 - 1.06 0 - 0.73

1 Diet designation: 65:00 = 65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gelatinized starch, 65:35 = 65 parts basal diets, 35 parts gelatinized starch, 100:00 = 100 parts basal diet, 0 part gelatinized starch.

2 Mean of the mean concentration of 3 tanks3 Range of values for individual fish4 Results from comparison of means using orthogonal contrasts with =0.05.

Estimated urinary glucose concentration (mM) and estimated urinary glucose

excretion rates (mmol/kg per h) were plotted against plasma glucose concentration and

presented in Figure 5.1 and 5.2 respectively. While there is significant variability, these

two figures show a relationship between plasma glucose and urinary glucose excretion

which is similar to what is observed in mammals (Khoushanpour 1976). The renal

threshold for glucose excretion in this experiment estimated using various methods

(Table 5.3) was estimated to be between 3.6 and 10.6 mM plasma glucose. Linear

regression suggested a renal threshold between 5.1 - 5.6 mM.

Figure 5.1 Urinary glucose concentration (mM) of rainbow trout fed the various

experimental diets as a function of plasma glucose concentration.

Urinary glucose concentration and corresponding plasma glucose values measured from

individual fish (39 observations). Diet designation are as follows: 65:00 = 65 parts basal

diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts gelatinized starch,

65:35 = 65 parts basal diets, 35 parts gelatinized starch, 100:00 = 100 parts basal diet, 0

part gelatinized starch.

Figure 5.2 Estimated (calculated) urinary glucose excretion (mmol/kg per h) of

rainbow trout fed the various experimental diets as a function of plasma glucose

concentration.

Estimated urinary glucose excretion rate (mmol/kg per h) calculated as follows: average

urine flow rate (L/kg per h) for fish of tank "y" x urine glucose concentration (mM) of

individual fish in tank "y". (39 observations). Diet designation are as follows: 65:00 =

65 parts basal diet, 0 part gelatinized starch, 65:20 = 65 parts basal diet, 20 parts

gelatinized starch, 65:35 = 65 parts basal diets, 35 parts gelatinized starch, 100:00 = 100

parts basal diet, 0 part gelatinized starch

Table 5.3 Renal glucose excretion threshold of rainbow trout as determined by

various approaches.

Approaches Threshold

(plasma glucose)

Minimum plasma glucose (observed) resulting in an urinary

glucose concentration > 0.3 mM

3.6 mM

Maximum plasma glucose (observed) resulting in an urinary

glucose concentration < 0.3 mM

10.6 mM

Linear regression on urinary glucose concentration1 5.1 mM

Linear regression on estimated urinary glucose excretion2 5.6 mM

1 Plasma glucose concentration giving an urinary glucose concentration = 0 mM based on the linear regression model presented in Figure 5.1

2 Plasma glucose concentration giving an urinary glucose excretion = 0 mmol/kg per h based on the linear regression model presented in Figure 5.2.

In this experiment, UFR and GFR were calculated from a measurement on a

"pool" of 4 fish per tank, which may not be totally appropriate for estimating individual

fish glucose excretion rates as used in Figure 5.2. The measure of UFR from a group of

four fish, instead of that on individual fish was, nonetheless, necessary to maintain

experimental conditions (good feed intake, low stress level) representative of those found

in the study described in Chapter Four.

Fish fed with high levels of digestible carbohydrate had high levels of blood

glucose and excreted very significant amounts of glucose in their urine. This excretion

could represent a major source of energy losses in fish fed high dietary levels of

digestible carbohydrate. From the average urinary excretion rates presented in Table 5.2,

one can attempt to roughly calculate glucose excretion of the fish fed the experimental

diets in the main feeding trial in Chapter Four of this thesis. Glucose excretion per fish

over the 84d experimental period can be estimated at 0.2, 0.4, 3.3 and 1.4 g for the fish

fed the 65:00, 65:20, 65:35 and 100:00 diets respectively. These estimates are only

tentative and may only be indicative of the order of magnitude of glucose excretion.

Glucose excretion rates were estimated from UFR and urine glucose concentration at a

single point in time and may not be totally representative of a 24h period. Fish used in

the present study were larger (269 g BW/fish) than fish used in the study presented in

Chapter Four (8.1 - 87.1 g BW/fish). If urine production is related to metabolic rate

(oxygen consumption), small fish should be expected to produce more urine per unit of

body weight.

From the detailed energy budget presented in Chapter Four (Tables 4.7 to 4.11),

approximately 60-100 kJ carbohydrate energy (equivalent to 3.5 - 6 g of glucose) is

missing for the fish fed the 65:20 diet and about 190 kJ (11 g of glucose) for the fish fed

the 65:35 diet. The calculated amount of glucose excreted in the urine (UE) represents,

therefore, approximately 30% of the unaccounted energy for the fish fed the 65:35 diet.

This could explain at least in part the apparent discrepancies in the energy budget of fish

fed diets high in carbohydrates, such as observed in studies described in the previous

chapters. Metabolites of glucose, such as lactate, could also be lost in the urine and

contribute to urinary energy loss. A large proportion (70%) of the discrepancy ("energy

gap") in the energy budget of the fish, nonetheless, still needs to be accounted for.

Branchial glucose excretion?

While not measured in this study, glucose excretion through the gills, a form of

non-fecal loss (ZE), may contribute to the apparent discrepancies in the energy of fish

fed high levels of digestible carbohydrates. A study using perfused head preparations has

shown that glucose may be excreted through the gills of rainbow trout (Furspan and Isaia

1983). This study showed that glucose excretion through the gills had a similar kinetics

to that observed for the kidney and that a similar glucose reabsorption system to that of

the nephron is involved (phlorizine sensitive, saturable transport mechanism). The

threshold for branchial excretion of glucose was found by that study to be around 2.5

mM and the reabsorption was completely saturated at about 55 mM of glucose in the

perfusing fluid. Glucose excretion rates obtained with the perfused head were about 10%

of the rate observed in the present investigation. While this model, like other perfusion,

ex vivo or in vitro models, may yield interesting information on glucose reabsorption /

excretion dynamics in the gills, great care should be taken when attempting to apply flux

obtained with this model to intact, live, unrestrained and fast growing fish. Given very

similar glucose reabsorption and excretion dynamics in the kidney and the gills of

rainbow trout, it may be speculated that the gills could also be a contributor to energy

loss by fish with elevated plasma glucose levels. The amount of glucose lost through the

gills may, however, be lower than what is lost through the urine. This speculation is

based on the low excretion rate obtained in the perfused head study of Furspan and Isaia

(1983) and on the assumption that only 20% of total PEG-4000 excretion was

attributable to branchial excretion in this study, compared to 80% for urinary excretion.

Branchial excretion of PEG-4000 could probably occur through the non-respiratory cells

(chloride cells) of the gills since inulin, a widely-used glomerular filtration marker, was

shown to be excreted by chloride cells (Isaia 1984). Chloride cells represent a small

proportion of the gills surface (Isaia 1984). These cells apparently possess a

tubulovesicular reticulum connected to the low pressure central venous sinus of the gills

and this system could constitute a mechanism of excretion similar to that of the nephron

(Isaia 1984).

Chapter SIX. GENERAL DISCUSSIONChapter SIX. GENERAL

DISCUSSION

The dietary model and pair-feeding protocol used in the studies presented herein

offered a simple, yet effective way of studying utilization of digestible carbohydrate by

rainbow trout. The work presented in this thesis is unique since it combines comparative

carcass analysis, digestibility, respirometry and estimation of urinary excretion of

glucose to quantify the various components of the energy budget of fish at increasing

digestible carbohydrate intakes.

In successive studies, it was confirmed that the energy from digestible

carbohydrate was poorly retained by rainbow trout. In a respirometry trial, it was

observed that fish fed the supplemental carbohydrate did not consume significantly more

oxygen that fish fed the basal diet alone, a finding which suggests that the digestible

carbohydrate was not aerobically metabolized. Excretion of glucose in the urine of fish

fed high levels of digestible carbohydrate was observed. This represented a significant

loss of energy and explained, at least in part, the apparent discrepancies in the energy

budget of fish fed supplemental carbohydrate in the studies presented here.

The various components of the energy budget accounted for approximately 90%

of the IE of the fish fed the 65:00 and 100:00 diets, leaving deviations (unquantified

energy losses) of about 10% (Table 4.11). The deviations, nonetheless, were increased

significantly by feeding the starch, indicating that other significant sources of energy loss

remain unindentified and unquantified. Urinary glucose excretion could only account for

30% of the unaccounted energy. Loss of glucose through the gills is a possibility since

both kidney and gills have a similar glucose reabsorption system. There is also a

possibility that a certain amount of lactate or other metabolites of glucose be lost through

the urine but this remains to be examined.

There was no difference in nitrogen retention or in the lipid content and fatty acid

profile (data not shown) of the carcass for fish fed the various levels of digestible

carbohydrate. The digestible carbohydrate, therefore, did not exert any negative

influence on the metabolism of amino acids and lipids.

The indirect calorimetry method used here (oxygen consumption) may be a

reasonably good method to estimate heat losses through aerobic metabolism (catabolism)

but does not allow the estimation of heat losses through anaerobic metabolism.

Estimation of heat production based on indirect calorimetry has been shown to

underestimate heat production under conditions in which anaerobic metabolism is

relatively important (Brafield and Llewellyn 1982). Catabolism of carbohydrate through

anaerobic processes (production of lactate, cycling through HMS) could involve loss of

heat that could not be accounted for by indirect calorimetry based on oxygen

consumption. The use of direct calorimetry would be desirable to verify this hypothesis

but this method is known to be insensitive and impractical for fish (Cho and Kaushik

1990).

Based on the evidence reviewed in Chapter Two (section 2.2.4), it can be

speculated that a significant source of energy loss could be through catabolism of glucose

to lactate in the intestinal tissue, followed by uptake of lactate by the periportal region of

the liver, synthesis of G-6-P through gluconeogenesis and cycling/degradation of the G-

6-P through the HMS. Significant loss of energy as well as the loss of one carbon atom

as CO2 would occur at every completion of a cycle through the HMS (Figure 2.1). This

could represent a futile cycle in which a portion of the digestible carbohydrate is

degraded. The lactate produce by the intestine and other tissues could also be excreted

by the animal resulting in a significant loss of energy.

The CO2 generated by the HMS would be confounded with the CO2 produced by

the TCA cycle. Production of CO2 at the HMS level would, therefore, introduce a bias in

substrate utilization calculations based on the respiratory quotient (RQ, CO2/O2). This

would increase the non-protein RQ values and would result in an overestimation of the

amount of carbohydrate (glucose) catabolized by the TCA cycle. The non-protein RQ of

rainbow trout fed high dietary levels of digestible carbohydrate has been determined by

Kaushik et al. (1989), Médale et al. (1991), Brauge (1994) and Médale et al. (1994). The

non-protein RQ calculated by these investigators are often high and suggested that a large

proportion (e.g. 30%) of the heat production of these fish is attributable to the (aerobic)

catabolism of carbohydrate. These results may appear in contradiction with the results

from the respirometry trial presented in this thesis. The generation of CO2 at the HMS

level (and wasting of a significant proportion of the carbohydrate-energy) could offer an

explanation for the difference in the conclusions reached in the studies of Médale et al.

(1991), Brauge (1994) and Médale et al. (1994) and those reached in the present thesis.

Since there is still a significant discrepancy (gap) in the energy budget presented

in this thesis, further studies on partitioning of energy from carbohydrates are warranted.

The quantification of the CO2 production of fish fed according to the dietary model used

in the present thesis would be interesting. The measurement of the flux through the

HMS would also be particularly interesting. Various methodological approaches have

been proposed by Katz and Wood (1960) to study the flux of glucose metabolites through

the HMS. These approaches involve the use of strategically-labelled glucose, such as 3,

4 14C glucose, and measurement of the extent of randomization of the label in metabolic

intermediates (G-6-P, trioses phosphate) or end-products (glycogen). In such a study,

however, the labelled glucose would have to be provided through the diet (not by

intravenous or intraperitoneal routes) since the intestinal mucosa may play a significant

role in the metabolism of glucose in fish (Harris 1993). In that respect, studies that have

looked at the utilization of intravenously injected (labelled) glucose by fish may not offer

an accurate picture of the glucose metabolism.

The suggestion that there is significant cycling of glucose metabolites through the

HMS raises some questions. Cycling of substrate through the HMS would result in the

production of NADPH whose electron would eventually have to be transferred to other

accepting molecules. The electron from NADPH can be transferred to NAD by

transhydrogenase or used in biosynthetic processes (fatty acid synthesis and elongation,

steroid synthesis). The electron transferred to NADH could be used to generate ATP

through oxidative phosphorylation. In the present thesis, oxygen consumption and fatty

acid profile of carcass lipid were unaffected by increasing the digestible carbohydrate

level, which may argue against a significant increase in the activity of the HMS. There

is, however, a possibility that the electron be accepted by a molecule which could,

subsequently, be excreted. This phenomenon has been observed in goldfish (Carassius

auratus) during anoxia (Mourik et al. 1982). Mitochondria of goldfish were shown to

decarboxylate a significant amount of pyruvate to acetaldehyde under anaerobic

conditions. After transport to the cytoplasm, acetaldehyde can act as an electron acceptor

for NADH, leading to the production of ethanol. Ethanol is known to easily permeate

membranes and diffuse into the surrounding water (Mourik et al. 1982).

From the results of the studies presented here, it is clear that no single ME or NE

value can be attributed to digestible carbohydrates. The amount of energy excreted in the

urine and recovered in tissues will depend on the intake of digestible carbohydrate. The

present studies also suggest that there is a threshold past which digestible carbohydrate

contributes nothing in terms of ME or NE. The NE value of the 14% digestible starch

used in our basal diet is not totally defined, but the good performance and feed efficiency

of the fish fed this diet as well as results from other studies (e.g. Hilton and Atkinson

1982) suggests that this small amount of carbohydrate can contribute useful energy to

rainbow trout. It might be interesting to conduct a similar experiment using a basal diet

without carbohydrate, and to increase allocation of digestible carbohydrate to determine

how much can be used efficiently or determine the ME and NE of digestible

carbohydrates as a function of intake.

The approach used in this thesis to determine the nutritive (energy) value of

carbohydrate for rainbow trout could be applied to other fish, such as the channel catfish

(Ictalurus punctatus), which are believed to be able to utilize carbohydrate more

efficiently than salmonids (NRC 1983, Wilson 1991, NRC 1993, Wilson 1994).

Practical diets for channel catfish are, in general, high in starch (>25-50%). Starch is

believed to be a significant energy source for the animal. These fish accept very well this

type of diet and attain good growth when fed with such diet in sufficient amounts (Page

and Andrew 1973, Mohsen and Lovell 1990, Li and Lovell 1992). This could, however,

have nothing to do with the actual nutritive value of the carbohydrate, but rather to the

ability of the fish to compensate for dilution of the diet with carbohydrates (and non-

nutritive materials) by consuming more diet. This is, perhaps, well illustrated by the

good growth but low feed efficiency of channel catfish fed practical diets containing high

levels of carbohydrates.

An analysis of the energy partitioning of channel catfish was performed by this

author using results from published studies (Page and Andrew 1973, Garling and Wilson

1976, Garling and Wilson 1977, Gatlin et al. 1986, Wilson and Poe 1987, Mohsen and

Lovell 1990) and the energy partitioning scheme used in this thesis. This analysis

suggests that, in reality, the ability of channel catfish to use dietary carbohydrate may not

differ much from that of rainbow trout (results not shown). This question should,

therefore, be addressed in a formal research trial. The use of a dietary model and

experimental protocol similar to that used in this thesis could help to determine

realistically the nutritive value of digestible carbohydrates for channel catfish and other

warmwater omnivorous species.

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