TED, ROLE OF HORMONES IN HEPATIC CARBOHYDRATE METABOLISM · 4.2.2 Hepatic glycogen accumulation in the starved diabetic rat 193 4.2.3 Properties of glycogen synthetase and phosphorylase

TED, ROLE OF HORMONES IN HEPATIC CARBOHYDRATE METABOLISM

A thesis submitted for the degree of

Doctor of Philosophy

in the

University of London

by

Patricia Denise Williams (nee Whitton)

Department of Biochemistry Imperial College of Science and Technology London, S.W.7 2AZ., U.K.

September, 1975

2

abstract

Net rates of glycogen accumulation (followed in sequential

liver samples) were measured in the perfused liver of 48h- starved

rats in the presence of glucose and gluconeogenic precursors. The

role of glucose as a carbon source of glycogen was assessed and found

to be minimal, gluconeogenic precursors beingthe major substrate of

glycogenesis. The activities of glycogen synthetase and phosphorylase

were assessed; their activities were related to the rate of glycogen

synthesis, and to the nature of the circulating substrates.

The achievement of normal net rates of glycogen accumulation,

in vitro, permitted the investigation of the role of hormones in glycogen

synthesis. Hormones of the posterior-pituitary gland,at "physiological"

concentrations,were found to cause glycogenolysis, prevent glycogen

synthesis and increase gluconeogenesis in the perfused liver. Although

the full mechanism of vasopressin action was not elucidated, increased

hepatic phosphorylase activity was found.

The role of insulin in hepatic carbohydrate metabolism was

assessed. An impairment in glycogen accumulation and response of glycogen

synthetase to substrates in the perfused liver of starved diabetic rats

was observed, which was restored by insulin, or glucose and fructose

treatment in vivo. Insulin did not, however, have any effect in vitro.

Glycogen accumulation and the response of glycogen synthetase and,

phosphorylase to substrates were impaired during perfusion of livers

from starved adrenalectomised rats. Restoration of net rates of

glycogenesis were observed after treatment in vivo with hydrocortisone,

or fructose, glucose and insulin, prior to perfusion, confirming previous

suggestions that the action of glucocorticoids is mediated by insulin.

However, no direct hepatic effect of insulin was obtained.

3

In conclusion, it would appear that glucose is not the major

precursor of hepatic glycogen (perhaps even in.the fed state), and

that adrenal corticosteroids and insulin exert crucial regulatory

effects on glycogen metabolism)which are not direct actions on the

liver. There are however, direct hepatic effects of vasopressin, whose

significance is not fully clear.

ACKNOWLEDGMENTS

I wish to express my sincere thanks to Dr. D. A. Hems for

his supervision, encouragement and discussions during the tenure of

the work reported in this thesis. I would also like to thank Professor

Sir Ernst Chain, F.R.S., and Dr. Anne Beloff -Chain for their

encouragement and interest in the project. \

My thanks are due to members of the Department, past and present,

(especially those who have resided in Rooms 5021'205) for many

stimulating discussions, andhelp. In particular I would like to thank,

Dr. I. Das for advice concerning the enzyme assays, and Mr. E. Taylor

and Mr. C. Harmon for excellent technical assistance. I am indebted

to Mr. D. Green for aid in the animal work, in breeding the rats and

help in the preparation of diabetic and adrenalectomised animals. The

amino acid analyses were kindly performed by Mr. C. Dykes.

My thanks are offered to Dr. M. Forsling (Department of Physiology,

Middlesex Hospital, London) who kindly carried out the bioassays of

vasopressin.

I am also indebted to Mrs. R. Sayer for the valuable "sun" time

she forfeited in order to type this thesis which must have seemed "double

-dutch" to her.

Finally, I would like to thank the Medical Research Council, U.K.,

for financing this project.

5

TO MY MOTHER, FATHER, HUSBAND AND MAX.

6

Investigating the mechanism of action of insulin on the glycogen

synthetase system is like peeling an onion. Not only may it bring

tears to the eyes, but after each successful step, one is left with

the layer beneath.

Wolff & Wolff (1964)

CO

Page

Abstract .2

Acknowledgements

Abbreviations and enzymenomenclature•

List of tables 16

List of figures

18

Chapter 1 : Introduction

21

Chapter 2 : Animals, Materials & Methods 47

Chapter 3 : Results • 73

Chapter 4 : Discussion 180

For details of each chapter see next page.

References 213

Page CHAPTER. ONE

INTRODUCTION

1.1 Glycogen metabolism in the liver 22

1.1.1 The characteristics of glycogen 22

1.1.2 The pathways of hepatic glucose metabolism 24

1.2 Hepatic glycogen synthetase and phosphorylase 27

1.2.1 General considerations 27

1.2.2 Hepatic glycogen synthetase 29

1.2.3 Hepatic glycogen phosphorylase 31

1.3 The circulating precursors of hepatic glycogen 33

1.4 The role of hormones in hepatic glycogen

35 metabolism.

1.4.1 Insulin and hepatic glycogen

35

1.4.2 Adrenal cortical steroids and hepatic glycogen 39

1.4.3 "Glycogenolytic" hormones and hepatic glycogen 42

1.5 Scope and aims of the present study

43

1.5.1 General considerations

43

1.5.2 The use of the perfused liver for metabolic

43 studies

CHAPTER TWO

ANIMALS, MATERIALS AND METHODS

2.1 The preparation of animals 49

2.2 Sourcesof materials 54

2.3 The technique of liver perfusion 56

2.3.1 Perfusion apparatus 56

2.3.2 Perfusion medium 59 ,

.2.3.3 Surgical procedure for liver perfusion 61

2.3.4 Sample preparation 63

9

2.4 The techniques used in intact animal

Page,

experiments 64

24.1 Measurement of net glycogen accumulation in vivo 64 '

2.4.2 Measurement of the enzymes of glycogen metabolism in response to hormones 65

2.5 Analytical methods ' 66

2.5.1 Glucose and glucose polymer determination 66

2.5.2 Carbohydrate metabolite determination 67

2.5.3 Determination of nitrogenous compounds 69

2.5.4 Assay for the enzymes glycogen synthetase and phosphorylase

CHAPTER THREE

RESULTS

3.1 The characteristics and control of hepatic looens -EIEMthesisilhe 8h- starvedrat

74

3.1.1 The validation of sequential sampling in the perfusion 74

3.1'.2 The role of glucose and gluconeogenic precursors in glycogen deposition in the perfused liver 77

3.1.3 The role of insulin and fatty acids in hepatic glycogen metabolism 188

3.1.4 Characteristics of glycogen synthetase and phosphorylase in the liver 90

3.1.5 Control of glycogen synthesis in the perfused liver of normal starved rats 97

3.1.6 Hepatic glycogen accumulation in the intact rat 99

3.2 Hepatic carbohydrate and fat metabolism in the "fed" rat 103

3.2.1 Glucose metabolism in the perfused liver of fed rats 103

3.2.2 Glycogen synthesis in the perfused liver of overnight-starved rats 105

10

Page:

3.2.3 Pattyacidsynthesis in the perfused liver of overnight-starved rats 107

.3.3 The actions of the hormones of the posterior-pituitary gland on hepatic glycogen metabolism 109

3.3.1 Stimulation of hepatic glycogen breakdown by (8 -arginine) -vasopressin and oxytocin 109

3.3.2 The role of vasopressin in hepatic glucose metabolism 115

3.3.3 The stimulation of hepatic gluconeogenesis by (8 -arginine) -vasopressin 117

3.3.4 The action of vasopressin and oxytocin on glycogen synthesis in the perfused liver and the intact rat 120

3.3.5 The effects of vasopressin and other glycogenolytic hormones on hepatic glycogen synthetase and phosphorylase in vivo 122

3.4 Hepatic glycogen metabolism in the starved streptozotocin-diabetic rat 128

3.4.1 Glycogen accumulation in the perfused liver from diabetic rats 128

3.4.2 Glycogen synthetase and phosphorylase activities in vivo and in the perfused liver of diabetic rats 133

3.4.3 Influence of glucose and fructose on the, activity of glycogen synthetase and phosphorylase in vivo 143

3.4.4 Hepatic glycogen accumulation in the intact diabetic rat 145

3.5 Hepatic carbohydrate metabolism in the starved adrenalectomised.rat 148

3.5.1 Glycogen accumulation In the perfused liver from adrenalectomised rats 148

.3.5.2 Glycogen synthetase and phosphorylase activities in vivo and in the perfused liver of adrenalectomised rats 153

3.6 Amino acid balance in the perfused liver 168

11

Page 3.6.1 Amino acid metabolism in fed, starved

and diabetic rat liver 168

3.6.2 Urea formation in the perfused liver 173

3.6.3 The role of the liver in amino acid utilisation 177

CHAPTER FOUR

DISCUSSION

4.1 Hepatic glycogen metabolism in the normal rat

4.1.1 The circulating precursors of hepatic glycogen 181

4.1.2 The role of glucokinase in glycogen accumulation 185

4.1.3 Control of hepatic glycogen synthesis

186

4.2 The role of insulin in hepatic carbohydrate metabolism

4.2.1 Insulin and hepatic glycogen metabolism in the normal starved rat 191

4.2.2 Hepatic glycogen accumulation in the starved diabetic rat 193

4.2.3 Properties of glycogen synthetase and phosphorylase in the perfused liver of diabetic rats 197

4.3-- The role of adrenocortical steroids in hepatic glycogen metabolism

4.3.1 Glucocorticoids and hepatic glycogen metabolism in the normal (starved) rat '201

4.3.2 Hepatic glycogen accumulation in the starved adrenalectomised rat 201

4.3.3 The characteristics of glycogen synthetase and phosphorylase in the perfused liver of adrenalectomised rats 205

4.4 The role of the hormones of the posterior- pituitary_gland in hepatic carbohydrate metabolism

12

Page

4.4.1 The posterior-pituitary gland hormones and the metabolism of liver carbohydrate 208

4.4.2 The mechanism of vasopressin action. 210

13

ABBREVIATIONS AND ENZYME NOMENCLATURE

Units and Physical Constants

Those used were as recommended in Biochem. J.

(1975) 145, 1

Chemicals

Abbreviations for amino acids are as in Biochem. J.

(1972) 126, 773

ADP .adenosine 5 -pyrophosphate

AMP adenosine 5

1 -phosphate

ATP adenosine 5' -triphosphate

Butyl PBD

cyclic AMP

5-(4-biphenyly1)-2(4-t-butylphenyl) -1 -oxa -3, 4- diazole

adenosine 3': 51 -cyclic

phosphate

cyclic GMP guanosine 3 : 5'-cyclic phosphate

EDTA ethylenediamine-tetra- acetic acid

K0104

potassium chlorate

KCN potassium cyanide

ICP potassium fluoride

K3Fe(CN)6 potassium ferricyanide

KOH potassium hydroxide

LiBr lithium bromide

MgC12 magnesium chloride

NaC1 sodium chloride

NAD nicotinamide-adenine dinucleotide

14

NADH2 nicotinamide-adenine

dinucleotide, reduced

NADP nicotinamide-adenine dinucleotide phosphate

RALPH.2 nicotinamide-adenine dinucleotide phosphate, reduced

NaF sodium fluoride

Nall2PO4

sodium dihydrogen orthophosphate

Na2BP04

disodium hydrogen orthophosphate

Pi orthophosphate

TCA trichloroacetic acid

Trig 2-amino-2-hydroxymethyl- -propane-1,3-diol

UDPG uridine diphosphoglucose

Other Abbreviations

Adx. adrenalectomised

intragastric

S.C. subcutaneous

15

Enzyme Nomenclature

The names of the enzymes are those recommended by a

I.U.P.A.C., and I.U.B., Commission, 1972. In a few instances,

trivial names are used and the Commission's recommendations are

included in brackets.

E.C. Number

Amyloglucosidase/Lysosomal tC-glucosidase 3.2.1.3. (Eko-1, 4-4-glucosidase)

Glucokinase 2.7.1.2.

Glucose oxidase 1.1.3.4.

Glucose 6-phosphatase 3.1.3.9.

Glucose 6-phosphate dehydrogenase 1.1.1.49.

Glycogen branching enzyme ; 2.4.1.18. Amylo-1, 4 .41, 6 glucosidase

Glycogen debranching enzymes ; Amylo-1, 6-glucosidase 4 oligo-1, 4 >1, 4 glucan transferase (4-oG- Glucanotransferase)

Glycogen phosphorylase

Glycogen phosphorylase kinase

Glycogen phosphorylase phosphatase

Glycogen synthetase (Glycogen synthase)

3.2.1.33. 2..4.1.25.

2.4.1.1.

2.7.1.38.

3.1.3.17.

2.4.1.11.

Glycogen synthetase kinase 2.7.1.37. (Protein kinase)

Glycogen synthetase phosphatase

Hexokinase 2.7.1.1.

Lactate dehydrogenase 1.1.1.27.

Peroxidase 1.11.1.7.

Phosphoglucomutase 2.7.5.1.

UDPG pyrophosphorylase 2.7.7.9. (Glucose 1-phosphate uridylytransferase)

16

LIST OF TABLES

Page

1. The effect of time of day on rat body weight and blood glucose concentration.

2. Seasonal variation in the growth of adrenalectomised rats.

Glycogen content and synthesis in the major lobes of perfused liver from 48h- starved rats.,

Glycogen synthesis and changes. in medium glucose during perfusion of livers from starved rats.

Calculated total synthesis of glucose during perfusion of liver from starved rats.

6. , Incorporation of 14

C from cg-14 C)-glucose (30mM) into glycogen of perfused livers from starved rats.

. 7a. Rates of net glycogen accumulation in perfusions with single substrates.

7b. Rates of net glycogen accumulation in perfusions with substrate conbinations.

8. Influence of added sodium oleate and insulin on glycogen synthesis in the perfused liver from starved rats.

The effect of different assay times on the activities of glycogen synthetase and phosphorylase.

10. The effect of variation in temperature and purification on the activities of hepatic glycogen synthetase and phosphorylase in the fed and starved rat.

11. Concentrations of enzymes and pathway intermediates during glycogen synthesis in the perfused liver of starved rats.

53

75

78

80

85

86

87

89

93

95

• i98

12. Glycogen synthesis in the perfused liver of overnight starved rats. 106

13. Effect of (8-arginine)-vasopressin on hepatic glycogen synthesis in vivo. 123

14. Effect of glycogenolytic hormones on, the activity of glycogen synthetase. 126

15. Glycogen accumulation during perfusion of livers from streptozotocin-diabetic rats. 130

16. Activities of glycogen synthetase and phosphorylase in intact starved normal and starved streptozotocin - diabetic rats. 134

17. Glycogen synthetase activity in the perfused liver of starved streptozotocin-diabetic rats. 135

18. Phosphorylase activity in the perfused liver of starved streptozotocin -diabetic rats. 137

19. Glycogen synthetase activity in the perfused liver of starved streptozotocin-diabetic rats. 141

20. Influence of glucose and fructose on glycogen synthetase and phosphorylase in intact starved normal and streptozotocin -diabetic rats. 144

21. Glycogen accumulation in intact starved normal and starved streptozotocin -diabetic rats. 146

22. Glycogen accumulation in the perfused liver from starved adrenalectomised rats. 150

23. Glycogen synthetase and phosphorylase activities in intact sham adrenalectomised and adrenalectomised rats. 155

24. Glycogen synthetase activity in the perfused liver from starved adrenalectomised rats. 156

25. Glycogen phosphorylase activity in the perfused liver from starved adrenalectomised rats. 158

26. The concentration of amino acids after perfusion in the presence of "normal" levels or no added amino acids. 171

27. Effect of added amino acids on urea production in the perfused liver. 174

28. Nitrogen balance in the perfused liver. 176

18

LIST OF FIGURES

Page

1. The pathway of liver glycogen metabolism. 25

2. Growth curve of streptozotocin and control injected rats. 49

Growth curve of adrenalectomised, sham-operated, normal on 0.9% Nacl and normal rats. 52

4. Apparatus for the perfusion of rat livers. 57

Rates of glycogen synthesis at various glucose concentrations in the perfused liver of starved rats. 79

6. Time course of glycogen synthesis in the perfused liver.. 82

7. Time course of glucose formation in the perfused liver. 83

8. Time courses of glycogen synthetase and phosphorylase 92 assays.

9. Rates of glycogen synthesis in vivo, at various blood glucose concentrations. 100

10. Time course of glycogen synthesis in vivo. 101

11. Glucose metabolism in the perfused liver of fed rats. 104

12. Influence of vasopressin on the time course of glucose output in the perfused liver of fed rats. 110

13. Dependence of the stimulation of hepatic glucose output on vasopressin concentration. 111

14. Effect of vasopressin on glycogen content of the perfused liver.

15. Influence of oxytobin on the time course of glucose output in the perfused liver of fed rats.

16. Effect of vasopressin on glucose metabolism in the perfused liver of fed rats.

17. Influence of vasopressin on gluconeogenesis or net glycogen accumulation in the perfused liver of starved rats.

-113

114

116

118

18. Effect of vasopressin on gluconeogenesis (endogenous and from added substrate)in the perfused liver of starved rats. 119

19

Page

19. Effect of (6 -lysine) -vasopressin and oxytocin on glycogen synthesis in the perfused liver of starved rats. 121

20. The time course of phosphorylase activation by glycogenolytic hormones. 124

21. Time course of net glycogen accumulation in normal and diabetic rats. 129

22. Relationship between net hepatic glycogen accumulation and glycogen synthetase and phosphorylase in diabetic rats. 139

23. Relationship between the changes in glycogen synthetase and phosphorylase during liver perfusion in diabetic rats. 142

24. Time course of restoration of net rates of glycogen deposition in the perfused liver from adrenalectomised rats. 152

25. Change in sensitivity of glycogen synthetase to substrates during net glycogen accumulation in perfused livers from adrenalectomised rats. 160

26. Response of phosphorylase activity during net rates of glycogen accumulation. 162

27. Relationship between the changes in glycogen synthetase and phosphorylase during liver perfusion in adrenalectomised rats. 164

28. Changes in combined response of both glycogen synthetase and phosphorylase during rates of net glycogen accumulation in the perfused liver from adrenalectomised rats. 166

29. Metabolism of amino acids during perfusion at high initial concentrations. 1169

30. The effect of four times "normal" concentrations of amino acids on urea production in the perfused liver. 175 •

CHAPTER ONE

INTRODUCTION

21

CHAPTER 1

INTRODUCTION

1.1 GLYCOGEN METABOLISM IN THE LIVER

1. The characteristics of glycogen

2. The pathways of hepatic glucose metabolism

1.2 HEPATIC GLYCOGEN SYNTHETASE AND GLYCOGEN PHOSPHORYLASE

1. General consi derat ions

2. Hepatic glycogen synthetase

Hepatic glycogen phosphorylase

1.3 !WE CIRCULATING PRECURSORS OF HEPATIC GLYCOGEN

1.4 THE ROLE OF HORMONES IN HEPATIC GLYCOGEN METABOLISM

1. Insulin and hepatic glycogen

2. Adrenal cortical steroids and hepatic glycogen

"Glycogenolytic" hormones and hepatic glycogen

1.5 SCOPE AND AIMS OF THE PRESENT STUDY

1. General considerations

2. The use of the perfused liver for metabolic studies

22

1.1 GLYCOGEN METABOLISM IN Thh LIVER

1.1.1 The characteristics of glycogen

Glycogen has been sometimes called animal starch. It is also

however, found in yeasts, algae and fungi and large amounts are found

in oysters and other shellfish. A similar polysaccharide has also been

found in the golden bantam sweetcorn. In higher animals, glycogen is

deposited in liver and muscle as a carbohydrate storage material,

available as an immediate source of energy.

The level of glycogen varies from tissue to tissue, the liver

being the main site of deposition where there may be up to 409/.4mol

of glycogen -glucose/g wet weight of liver, which can be rapidly depleted

by hormones or starvation. In skeletal muscle the level of glycogen is

only about 251Lmol of glyeogen-glucose/g which is not depleted on

starvation, but is decreased by exercise and hormones, especially

adrenalin. Glycogen is a glycolytic fuel in muscle, providing lactate,

but in the liver, due to, the presence of glucose 6 -phosphatase it is

also degraded to glucose. This glucose is then released into the

circulation to be utilised by other tissues especially skeletal muscle

and the brain. The above considerations show the importance of hepatic

glycogen and why the levels must be tightly controlled by hormones and

the blood glucose level.

Glycogen is a polymer in which glucose residues are cc-1, 4—

linked with0C-1, 6 branchpoints. A number of structures have been

proposed from available chemical and enzymatic data, e.g., the one

proposed by Whelan (1971).

It is widely accepted that the glycogen molecule is a compact

multi-branched structure, with an average chain length ofi2-)4glucose

residues. Due to the high degree of branching, about 10 per cent of

the glucose units are situated at non-reducing termini available for

degradation, enabling rapid mobilisation. The branching also confer

a high degree of solubility, whereas the large molecular weight exerts

a small osmotic pressure in comparison to the same amount of glucose in

a free form.

Several pioneering observations, which have been crucial in

the progress of biochemistry, may be cited for work in the glycogen

field (see Ryman & Whelan, 1971). Thus glycogen was the first polymer

to be synthesised in vitro, and the first for which it was recognised

that nucleoside diphosphate sugar and a primer were necessary for

synthesis. It is one of the few branched polysaccharides whose

molecular structure has been studied extensively and has been used as

a model for the enzymatic determinations of polymer structure..

Glycogen synthetase and glycogen phosphorylase (the enzymes involved

in the metabolism of glycogen) were the first enzymes shown to be

regulated by phosphorylation and dephosphorylation of the protein, a

phenomenon which now appears to be implicated in the control of a number

of enzymes. The adenine nucleotide, cyclic AMP, was discovered when

its role in the control of phosphorylase activity was recognised.

These and other "firsts" illustrate the contribution that the study

of glycogen has made to a number of different aspects of biochemistry.

1.1.2 The pathways of hepatic glucose metabolism

The pathway of glycogen metabolism in the liver is summarised in

Fig.1. Glycogen synthesis from glucose occurs by the successive

action of glucokinase (1), phosphoglucomutase (2), MPG -

pyrophosphorylase (3), and glycogen synthetase and branching enzyme'

(4 and 5). The phosphorylytic degradation of the polysaccharide

involves the co-ordinated operation of phosphorylase and debranching

enzyme (6 and 7); glucose 1-phosphate formed by phosphorolysis,

is converted to glucose by the successive action of phosphoglucomutase

(2) and glucose 6 -phosphates° (8). Glycogen degradation can also

occur by hydrolysis under the action of lysosomaloC-glucosidase

(Jeffrey et al., 1970; Lejeune et al., 1963). The role of this

mechanism in the control of physiological degradation of glycogen is

still uncertain.

The liver contains three enzymes capable of phosphorylating glucose,

namely hexokinase, glucokinase and glucose 6-phosphatase. Hexokinase

has a very high affinity for glucose (Km of rat liver enzyme about

3 x 10 -5M; Vinuela et al., 1963; Walker, 1963) and is thus always

saturated in vivo (glucose concentration 5 to 10mM). This enzyme

cannot therefore, be affected by variations in the blood suger level.

Glucokinase is however, an enzyme which can convert glucose into glucose

6-phosphate at a rapid rate. It displays a Km for glucose of 10-40 mM

(Vinuela et al., 1963; Walker, 1963) and this activity is thus highly

responsive to changes in glucose. Microsomal glucose 6-phosphatase,

in addition to its hydrolytic role, can catalyse phosphotransferase

reactions (see Nordlie, 1968 for review). The Km for glucose is 80 mM

and so it would appear that glucose phosphorylation by this enzyme

LUCOSE 1-P

U D PG

PP i

UD P

(GLUCOSE)

[1, 4--)1,6

(GLUCOS)n+1

GLYCOGEN

UTP

GLUCOSE 6-P

Fig. 1

(1) (2)

(3) (4) (5) (6) (7)

(8)

The pathway of liver glycogen metabolism

Glucokinase

Phosphoglucomutase

UPG — pyrophosphorylase

Glycogen synthetase

Branching enzyme

Phosphorylase

Debranching enzyme

Glucose 6—phosphatase

•

26

would not occur except at exceptionally high glucose concentrations;

this action has been invoked in diabetic animals (Friedmann et al.,

1967) which have a very low level of glucokinase (Vinuela et al.,

1963). As will be seen, the present experiments show there is no

requirement to invoke a role for this enzyme in glucose phosphorylation,

in diabetes or in any other situation.

1.2 HEPATIC GLYCOGEN SYNTHETASE& GLYCOGEN PHOSPHORYLASE


Glycogen synthetase and the branching enzyme (amylo

1, 4-41, 6 glucosidase) are involved in glycogen synthesis, and

phosphorylase and the debranching enzymes (olig0-1,4-)1, 4 glucan

transferase and amylo-1, 6-glucosidase) in glycogen breakdown.

Although the branching and debranching enzymes are important in

determining the structure of glycogen they do not appear to be rate

controlling (Birch et al., 1974; Ryman & Whelan, 1971). Glycogen

ynthetase and phosphorylase are however, rate-limiting for synthesis

and degradation respectively.

Glycogen synthetase catalyses the formation of 0C -1,

4-glucosyl bonds on the outer branches of glycogen utilising UDP -

glucose as the glucosyl donor:

DTP glucose + glycogen ------41111° + glycogen

(n glucosyl units) (n + 1 glucosyl units)

Salsas and Larner (1975) have shown that muscle glycogen synthetase

is able to use free glucose as the glucosyl acceptor, although the

presence of very high glucose concentrations were necessary. It may

therefore be unlikely that this occurs in vivo.

28

The discovery of glycogen synthetase in liver by Leloir and Cardini

(1957) ended an era in which it was thought that glycogen synthesis,

as well as degradation,,was catalysed by phosphorylase. The

equilibrium of the above reaction favours the formation of glycogen

and so the reaction is irreversible in vivo. Liver glycogen

synthetase is bound to liver glycogen (Leloir & Goldemberg, 1960)

which is found associated with the smooth endoplasmic reticulum (Pbrter

& Bruni, 1959). The enzyme is located in both the microsomal pellet

and supernatant, although greater activity is found in the former

fraction (Maddaiah & Madsen, 1968). Unlike phosphorylase however,

the distribution of glycogen synthetase does not appear to be influenced

when hepatic glyCogen content is changed by diet (Maddaiah & Madsen,

1968).

Glycogen phosphorylase catalyses the transfer of glucosyl units

from the non-reducing ends of the polysaccharide to inorganic phosphate:

Glycogen + + Glucose 1-P

(n glucosyl units) . glucosyl units)

The equilibrium of the reaction is reached when the ratio of

inorganic phosphate to glucose 1-phosphate is about 3 at pH 7 (Cori

et al., 1940 & 1943). Although this reaction could therefore be

reversible the reaction is primarily degradative in vivo due to the

high intracellular concentration of inorganic phosphate and low level

of glucose 1-phosphate. Subcellular fractionation of rat liver

suggests that phosphorylase is bound to liver glycogen in the well-fed.

state and sediments with the microsomal fraction. However, when

glycogen levels are depleted the enzyme is found in the supernatant

fraction (Maddaiah & Madsen, 19663 Tata, 1964).

29

1.2.2. Hepaticfllycoen synthetase

Two forms of glycogen synthetase have been isolated from

the liver, the "a" or "I" form which is dephosphorylated and the

"b" or "D" form which is phosphorylated (Bishop & Larner, 1969;

Eiznkuri & Larner 1964; Mersmann & Segal 1967). The possibility

of a third form of glycogen synthetase in muscle at least, has

been proposed by Rosell-Perez (Hildalgo & Rosell-Perez, 1971).

The "I" and "D" (i.e., independent and dependent on glucose

6-phosphate for activity) nomenclature is derived from the muscle

enzyme but as will be seen later does not strictly apply to the

liver enzyme. The two forms of the enzyme are interconverted by

a kinase requiring Mg and ATP (D6Uulf & Hers, 1968a) which is

irhibited by high ATP and high Mg2+ (Bishop & Larner, 1967 & 1969)

and activated by cyclic AMP (Bishop & Lamer, 1969; Glinsmann &

Bern, 1969; Jefferson et al., 1968) and a phosphatase (two forms of

this enzyme have been postulated by Bishop, 1970) which is Mg2.+

dependent (Hizukuri & Lamer, 1964), activated by glucose (DeWulf

& Hers, 1967a& 1968b) and inhibited by high concentrations of

glycogen (Hers et al., 1970) physiological concentrations of ATP

(Gilboe & Nuttall, 1973 & 1974; Gold, 1970a) and phosphorylase "a"

(Stalmans et al., 1971 & 1974a).

The two forms of glycogen synthetaSe are independent and

dependent on glucose 6-phosphate for activity (respectively) when

measured at saturating concentrations of MPG (Hornbrook et al.,

1966) and the nomenclature "In and "D" was based on this concept.

30

It is now apparent, however, that synthetase "I" is stimulated

by glucose 6-P when assayed in the presence of physiological

concentrations of DFDPG and is thus not independent Of the sugar

nucleotide for activity (Hornbrook et al., 1966; Mersmann & Segal,

1967) and that under these conditions the "D" form of the enzyme

is virtually insensitive to glucose 6-phosphate in the physiological

range of concentrations. The role of glucose 6-phosphate in the

regulation of liver synthetase has been doubted also by DeVUlf et

al., (1968)2who favour a major regulatory function for Pi. In

mouse liver, a physiological concentration of Pi (5)4mol/g)

stimulates the "I" form and overcomes inhibition due to 3mM ATP,

but inhibits the stimulation of the "D" form by glucose 6-P and

is unable to overcome the nucleotide inhibition of this form.

Therefore, at the concentrations of hepatic UDPG, ATP and Pi observed

in vivo, there is no stimulatory effect of glucose 6-P on the "D"

form but up to 60% stimulation of the "I" form. Since the "I" and

"D" nomenclature as originally defined for muscle does not apply

to the liver enzyme, and "b" will be used henceforth for the

"active" and "less active" forms of glycogen synthetase (see DeWulf

et al., 1968; Hers et al., 1970).

In view of the complexity of allosteric regulation of glycogen

synthetase in vitro, it is difficult to assess the role of

metabolites in modulating synthetase activity in vivo. In general,

it appears that the combined effect of inhibitory metabolites (ATP,

ADP, Pi) and stimulatory effectors (glucose 6-P, Pi, citrate) is to

maintain synthetase "a" activity and inhibit synthetase "b" in vivo,

in conditions conducive to glycogen deposition.

31

The role of hormones in the control of glycogen synthetase

will be discussed later (see Section 1.4; for reviews see Lamer &

Villar-Palasi 1971; Ryman & Whelan, 1971; Soderling & Park, 1974).

1.2.3 Hepatic glycogen phospho lase .

Two forms of phosphorylase exist in the liver, the "a" or

phosphorylated form and the "b" or dephosphorylated form (Sutherland

& Wosilait, 1956). They are interconverted by phosphorylase phosphatase

which is activated by glucose (Stalmans et al.,/19790 4:nd ATP (Merlevede

et al., 1969) and inhibited by AMP (Wosilait & Sutherland 1956), and

phosphorylase kinase which is activated by cyclic AMP (Rail & Sutherland

1958). There is some evidence to suggest that the phosphatase exists

in two forms (Merlevede et al., 1969)as is the case with the muscle

enzyme. The kinase has not been purified from liver. •

In muscle the "a" form is active in the absence of other factors

but the "b" requires AMP. The situation is not as clear in the case of

the liver enzyme for although "a" is intrinsically active, 1mM AMP

increases its activity by about 15 to 40% (dog liver: Sutherland &

Wosilait, 1956; rabbit liver: Wolf et al., 1970). In the absence of

AMP, phosphoi:ylase "b" was thought to be virtually inactive until the

recent work of Tan and. Nuttall (1974) who demonstrated that the rat

liver enzyme was active in the absence of the nucleotide. The

reported values for activity of phosphorylase "b" in the presence of

AMP are variable: 15% of total phosphorylase activity for dog liver

enzyme (Wosilait & Sutherland, 1956); 0.2% for pig and rabbit (Appleman

et al., 1966); up to 25% for mouse liver (DeWulf, 1971) and 80% for

rat liver (Tan & Nuttall, 1974).

The situation, therefore, as regards the activities of phosphorylase

"a" and "b" is not clear, although it is likely that in vivo,

phosphorylase "a" is the "active" form and that "b" is converted to

nan when an increase in phosphorylase activity is required. Such

a change is clearly established in response to hormones or hypoxia

for example; the roles of hormones and cyclic AMP on the enzyme

glycogen phosphorylase will be discussed later (see Section 1.4 and

for review see Fischer et al., 1970; Soderling & Park, 1974).

33

1.3 THE CIRCULATING PRECURSORS OF HEPATIC GLYCOGEN

The circulatory precursors which provide the carbon atoms of

hepatic glycogen have not been fully identified. Synthesis and

de gradation of hepatic glycogen is closely related to the intrinsic

hepatic homeostatic control of blood glucose concentration. It

has been shown (Soskin, 1938 & 1941) that when circulatory glucose

levels are high there is glucose uptake by the liver and that during

hypoglycaemia glucose is released by the liver.

The influence of increasing glucose levels has been widely

studied and in general it has been found that glycogen synthesis is

enhanced in a dose-dependent manner. This has been shown in a variety

of systems: liver slices (Ballard & Oliver, 1964; Cahill et al., 1958),

perfused livers (Buschiazzo et al., 1970; Glinsmann et al., 1970; Haft

1967; Rudman & Herrera, 1968; Sokal et al., 1958) and in vivo

(Madison, 1969). Based on the properties of glucokinase and glycogen

synthetase the following was proposed (Leloir, 1964 & 1967; Steiner

1964): an increase in glucose concentration is expected to accelerate

glucose phosphorylation and lead to an increase in the intrahepatic

concentration of glucose 6-phosphate; stimulated by this metabolite,

glycogen synthetase converts the glucose moiety of UDPG into glycogen.

An increase in glycogen accumulation could also occur because of the

known activation of hepatic glycogen synthetase phosphatase (DeWUlf &

Hers, 1967a & 1968b) and phosphorylase phosphatase (Stalmans et al.,1970

• 1974a)by glucose.

34

However, in the starved-refed state the role of glucose as a

glycogen precursor is likely to be diminished due to the reduced

levels of glucokinase on starvation (Salas et al., 1963; Walker &

Rao, 1964). When material other than glucose is administered after

starvation, hepatic glycogen deposition (Hornbrook et al., 1965 &

1966; Winternitz et al., 1957) is likely to be due to gluconeogenesis

(defined as the synthesis of glucose in a monomer or polymer form).

If, however glucose is ingested after starvation, the accumulation

of hepatic glycogen could be due to continued gluconeogenesis or

hepatic uptake of circulating glucose. From experiments in vivo

with 140-labelled precursors, Olavarria et al.,(1968) suggested that

hepatic glycogen synthesis in starved-refed rats,- even when they

receive glucose, is mainly a result of gluconeogenesis, at least

initially

One of the objectives of this work was to clarify which were

the circulatory precursors of hepatic glycogen in both starved and

fed animals, and determine the role of glucose in glycogen

accumulation.

35

1.4 THE ROLE OF HORMONES IN HEPATIC GLYCOGEN METABOLISM

Perhaps the most outstanding and least clarified aspect of

glycogen metabolism is that of the role of hormones in glycogen

metabolism, where many conflicting data exist, especially with respect

to insulin (for reviews see Nuttall, 1972; Pilkis & Park, 1974) and

adrenal cortical steroids (for review see Landau,l965). This could

be due to the fact that the hormones have been tested in isolated

liver preparations under suboptimal conditions, e.g., when net glycogen

synthesis has not been observed.

1.4.1 Insulin and hepatic glycogen

One experimental approach used to study the role of insulin

in metabolism is to make animals deficient in the hormone (diabetic)

and compare their carbohydrate metabolism with that of matched normal

animals. In diabetic animals there is an impairment of extrahepatic

glucose utilisation. Their low hepatic glycogen concentrations

compared with-those of the normal fed animals (Friedmann et al., 1963),

combined with a high blood glucose (as a result of increased

gluconeogenesis: Exton et al., 1972b& 1973a; Friedmann et al., 1965;

Renold et al., 1953), point to a major impairment in the glycogen-

synthesising mechanism. Loss of control of glycogen synthetase and

phosphorylase by glucose has been observed in the perfused liver of

diabetic rats (Miller et al., 1973) which correlates with the reported

loss of glycogen synthetase - activating system (Bishop, 1970; Gold,

1970b; Nichols & Goldberg, 1972).

36

This loss has not been confirmed by DeWulf (1971) who found that

glycogen synthetase phosphatase was normal in diabetic animals. The

diabetic animal has increased activity of hepatic glycogen synthetase

"b" (Kreutner & Goldberg, 1967; Steiner et al., 1961)'and an increase

in total activity (Steiner et al., 1961).

A number of studies using 14C-labelled substrates e.g., lactate

or alanine, have shown an impaired incorporation of 14C into hepatic

glycogen, although glucose becomes highly labelled (Exton et al.,

1972b& 1973a; Friedmann et al.,111Ar)Only on administration of

insulin in vivo does glycogen become labelled, in preference to

glucose (Exton et al., 1972b& 1973a) and an increase in activity of

glycogen synthetase"a"occur (Gold, 1970b; Kreutner & Goldberg, 1967;

Miller & Larner, 1973; Steiner et al., 1961 & 1964).

Although insulin appears necessary in diabetic animals for the

restoration of net glycogen accumulation (Steiner & King, 1964) and

of the activity of glycogen synthetasej several studies in vivo

suggest that synthesis can occur in insulin-deficient animals, without

administration of the hormone. Hornbrook (1970) and Friedmann et al.,

(1963 & 1967)_have reported normal rates of glycogen accumulation

although the eventual levels of glycogen stored were less than in the

normal animal and increased after insulin administration (Friedmann

et al., 1963 & 1967). They proposed that synthesis was not impaired

in the liver but that it was the capacity for glycogen storage which

was affected in diabetes.

Study of the role of insulin in vivo in normal animals has been

complicated by the role of glucose in hepatic glycogen synthesis

(Buschiazzo et al., 1970), glycogen synthetase (DeWulf & Hers, 1967a

& 1968b) and phosphorylase (StaImans.et al., 1970 & 1974a).

37

It has been proposed that glucose and not insulin, is the important

factor in the activation of glycogen synthesis (DeWulf & Hers, 1967a)

and glycogen synthetase (DeWulf,1971), and inhibition of phosphorylase

activity (StaImans et al., 1974a). This has been shown in perfusion

studies where the increase in glycogen synthetase and decrease in

phosphorylase activities due to glucose, were unaffected by the

presence of insulin (Glinsmann et al., 1970). Miller and Larner (1973)

have however, shown a direct rapid effect of insulin on glycogen

synthetase in perfused livers, and Hostmamk (1973) has shown in liver

perfusion that this effect is not mediated by or dependent on glucose.

The most consistent short-term hepatic action of insulin

observed in vitro has been to antagonise the effects of glucagon

and adrenalin (Exton et al., 1970 for review; Exton et al., 1973a;

Glinsmann & Mortimore, 1968; Hostmark, 1973). Under these conditions

a decrease in elevated cyclic AMP is observed, which is not apparent

when insulin is added alone. It has been suggested that this lack

of fall in cyclic AMP is due to the presence of a small metabolically

active pool upon which insulin acts (for reviews see Pilkis & Park,

1974; Walaas et al., 1974).

The discovery that insulin activated glycogen synthetase (in

muscle) led to the proposal of two rapidly interconvertible forms

of the enzyme (Villar-Palasi & Larner, 1960). Since then assay

systems have been developed which differentiate between these two

forms and have shown clearly that in intact animals, insulin causes

a conversion of the hepatic "b" (phosphorylated) form to the "a" (non-

phosphorylated) form.

38

Such activation could be a direct effect of insulin (if glucagon

is also present) mediated by prevention of both the formation and

action of cyclic AMP (Glinsmann & Mortimore, 1968). Whether there

is a separate "second messenger," different from cyclic AMP,

such as cyclic GMP (proposed by Illiano et al., 1973), or whether all

insulin effects occur via a decrease in cyclic AMP is unclear. It

is also uncertain whether insulin acts via the activation of glycogen

synthetase phosphatase (Bishop et al., 1970 & 1971; Gold, 1970b), or

. inhibition of glycogen synthetase kinase (both in the absence and

presence of glucagon: Miller & Lamer, 1973) and whether it has any

effect on glycogen phosphorylase.

As can be seen from the above there is still controversy over

the role and mechanik of insulin action in hepatic glycogen metabolism.

One group of the experiments reported here was designed to clarify this

issue.

39

1.4.2 Adrenal cort acsteroids a lauatisgia

The role of glucocorticoids in hepatic glycogen metabolism,

like that of insulin, is somewhat unclear. Although it has long

been known that the hypoglycaemia that kills adrenalectomised animals

upon prolonged starvation is due to an ultimate failure of

gluconeogenesis (Long et al., 1940), it is not clear how adrenal

steroids act on gluconeogenesis. It is known that (in intact animals)

they enhance the peripheral breakdown of proteins and stimulate the

uptake of Amino acids by the liver but their influence in isolated

liver preparations on conversion of amino acids to glucose is variable.

Friedmann et al., (1965) found that adrenalectomy did not impair

gluconeogenesis from pyruvate or alanine'inthe starved animal, unlike

Eisenstein et al., (1966) who found an impairment from alanine, which

was corrected by dexamethasome in vivo or in vitro. Exton and co.

workers also found a decrease in gluconeogenesis from lactate in the

starved adrenalectomised (1965) and diabetic-adrenalectomised rat

(1970 & 1973b) which was restored by treatment in vivo or in vitro

(1973b). Adrenalectomy curtails the stimulation of gluconeogenesis by

glucagon in the perfused liver (Exton et al., 1972a;Friedmann et al.,

1967); the glucagon response returns on treatment with glucocorticoids

in vivo. Although addition of steroids alone in vitro has no rapid

effect on gluconeogenesis (in the absence of glucagon), restoration

of the glucagon response is seen with dexamethasome (Friedmann et al.,

1967).

Although hepatic gluconeogenesis appears to be little altered

by steroids in vitro, hepatic glycogen synthesis is low after

adrenalectomy (Friedmann et al., 1965) and is not responsive to hormone

40

added in vitro (diabetic-adrenalectomised rats, Exton et al., 197313);

treatment in vivo for 2-3h being required for glycogen accumulation to be

restored (Hornbrook et al., 1966; Nichols & Goldberg, 1972). However,

it has been reported that hepatic glycogen synthesis is not

significantly impaired in vivo (Friedmann et al., 1967; Kreutner &

Goldberg, 1967) although a two hour lag in synthesis was observed

which was shortened by prior steroid treatment in vivo (Kreutner &

Goldberg, 1967). The rate of glycogenesis wasas rapid as in the normal

animal but "plateaued" at a lower level, indicating that steroids may

be involved in the amount of glycogen stored.

In the fasted•adrenalectomised rat most of hepatic glycogen

synthetase is in the inactive "b" form (Glinsmann et al., 1970;

Homnbrook et al., 1966; Mersmann & Segal, 1969) due to the low

activity of glycogen synthetase activating system; it is restored

after 2-3h hydrocortisone treatment in vivo (Gruhner & Segal, 1970;

Mersmann &.Segal, 1969). There does not however, appear to be such

an impairment in the fed adrenalectomised animal where the conversion

of glycogen synthetase "b" to "a" can be activated by glucose in vitro

(Glinsmann et al., 1970; Miller et al., 1973). The response of glycogen

phosphorylase to glucose is also impaired, in the starved adrenalect-

omised rat (Miller et al., 1973). These differences, between fed and

starved animals, could implicate insulin in steroid action (since

insulin levels are higher in fed animals). Although the lack of adrenal

corticosteroids results in no change in the levels of"activeu hepatic

glycogen phosphorylase, the levels of inactive enzyme are greatly

diminished; restoration of the enzyme activity and hyperglycaemic

effects of adrenalin and cyclic AMC' being obtained with steroid

replacement (Schaeffer et al., 1969).

Studies on the role of steroids in the normal animal have been

extensive (for review see Landau 1965), their administration in

vivo leading to an increase in blood glucose concentration, hepatic

glycogen and glycogen synthetase "a", and decrease in phosphorylase

activity (DeWulf & Hers, 1967b & 1968b; Hornbrook et al., 1966 & 1970).

There does however, seem to be a time lag (shorter than in

adrenalectomised animals) before a response is observed. The effects

of steroids on glycogen accumulation in vitro are not clear.

The restoration of hepatic glycogen accumulation and increase

in glycogen synthetase "a" (associated with an increase in glycogen

synthetase phosphatase) seen in adrenalectomised animals treated with

hydrocortisone (Nichols & Goldberg, 1972) is not observed within 2h

in adrenalectomised-diabetic rats (Exton et al., 1973b; Nichols &

Goldberg, 1972). Restoration of glycogen synthetase "a" activity

is however, seen after 2-4 min of insulin treatment (Nichols &

Goldberg, 1972). It was proposed (Hornbrook, 1970; Kreutner & Goldberg,

1967; Nichols & Goldberg, 1972) that the ability of steroids to induce

glycogen deposition is largely dependent upon insulin release. Insulin

secretion is reduced to 60X, by adxenalectomy OPIalaisse et al., 1967) and it

is thought that steroids increase the sensitivity of insulin secretion

to glucose.

It is not clear therefore, whether adrenal corticosteroids exert

direct effects on hepatic glycogen metabolism or whether their action

(in vivo) is mediated through insulin or glucose. These aspects were

investigated in the experiments to be described.

42

1.4.3 "Gle°n°1-bicithor icirc°ge

an

n

The above two hormones (insulin and glucocorticoids) are

generally considered to be involved in the control of glycogen

accumulation. Glycogen breakdown is however, as important a process,

since it is the balance between the two which governs the actual

amount of glycogen stored. The two main hormones known to cause

glycogenolysis are glucagon and adrenalin (Exton et al., 1970 for

review), although the importance of the latter hormone in hepatic,

glycogen breakdown in rodents is thought to be minimal (Exton & Park,

'1968; Sokal et al., 1964).

Additional hormones can cause glycogenolysis. For example,

early work demonstrated that extracts of the neurohypophysis could

cause hyperglycaemia (Clark, 1928; Claude & Baudouin, 1912) and

prevent the hypoglycaemia due to insulin (Burn, 1923) perhaps as a

result of breakdown of liver glycogen (Clark, 1928; Imrie, 1929; Stehle,

1950). When purer preparations of hormones became available

vasopressin was shown to bring about an increase in blood glucose in

mammals (Bergen et al., 1960; Cash & Kaplan, 1964; Schillinger et al.,

1972), possibly through hepatic glycogenolysis (Bergen et al., 1960;

Heidenreich et al., 1963). Evidence showing that this hyperglycaemia

was due to hepatic glycogenolysis was obtained by Heidenreich et al.,

(1963) in liver slices and Vaisler ( 1965a ) in perfased liver, who

found that oxytocin displayed a similar action, although the response

was smaller and of a shorter duration that vasopressin (Vaisler, 1965b).

The significance of these results has been uncertain, since the

concentrations of the hormones were usually higher than those which may

occur "physiologically". One portion of the investigation reported

here was designed to clarify the roles of vasopressin and oxytocin in

liver carbohydrate metabolism.

43

1.5 SCOPE AND AIMS OF THE PRESENT STUDY


From the above, it is obvious that the role of hormones in

particular in hepatic glycogen metabolism is unclear. As has been

stated, most of the work has been carried out in systems which have

not exhibited the rates of glysogen accumulation which are known to

occur in vivo.

The aim of this research was to obtain conditions in the

perfused liver which are suitable for maximal net rates of glycogenesis,

comparable to the in vivo rates, and to study the role of hormones in

this state. In parallel with measurements of glycogen and substrates,

the enzymes of glycogenesis and glycogenolysis have been measured under

conditions of maximal and sub-maximal glycogen accumulation. Glycogen

breakdown is as important in the liver as glycogen synthesis and the

role of hormones in this aspect of glycogen metabolism has also been

investigated, with a similar group of measurements.

1.5.2 The use of the perfused liver for metabolic studies

There are a number of different experimental techniques available

for metabolic studies. They may be divided into two groups: those

using the intact animal and those involving isolated tissue preparations.

The use of the intact animal for metabolic studies of a

particular tissue has a number of disadvantages. It is difficult to

produce a specific change in one tissue without stimulating parallel or

compensatory changes in others, or without initiating nervous or

hormonal discharges which could produce marked metabolic changes in the

tissue under investigation.

Thus metabolic studies in the whole animal do not yield insights

into the specific or primary sites of action of circulating hormones

or substrates, but may be used for obtaining confirmatory evidence

for theories of metabolic control.

There are four main types of isolated tissue preparation:-

(a) homogenates, (b) cells (c) pieces or slices and (d) organ perfusion.

Metabolic studies involving the use of homogenates are primarily

utilised for the assay of various tissue constituents including

enzymes. The use of tissue slices, on the other hand, has been a

very productive technique, a large part of basic metabolism (especially

the elucidation of pathways) being based on data obtained by this

method. Liver slices do however, have a number of disadvantages in

the study of metabolic control. They lose sugar phosphates,

nucleotides, proteins and ions and are subject to variable diffusion

properties. The use of hepatocytes in metabolic investigations has

an advantage compared with liver slices, in that metabolic functions

(e.g., gluconeogenesis) are maintained. However, until recently

their response to hormones did not appear to be as sensitive as the

perfused live.

Metabolic studies have been extensively carried out using the

perfused liver. It has a number of important advantages over the

techniques described above in that it is the most "physiological"

system, it involves the least disruptive procedures during the

preparation, and the perfusate may be easily controlled and monitored.

Per-haps however, the most important aspect of liver perfusion is that

the role of hormones may be evaluated, especially to elucidate whether

there is direct or indirect hepatic action.

45

Although this could be tested in slice or cell systems they have

the disadvantages given above with the additional factor that

hepatocyte studies tend to isolate particular cell types neglecting

the role of non-parenchymal cells, and the circulation within the

liver.

In view of considerations such as these, the liver perfusion

technique was employed in the present study.

CHAPTER TWO

ANIMALS, MATKRIALS & METHODS

47

CHAPTER 2

ANIMALS, MATERIALS AND METHODS

2.1 THE PREPARATION OF ANIMALS

2.2 SOURCES OF MATERIALS

2.3 THE TECHNIQUE OF LIVER PERFUSION.

1. Perfusion apparatus

2. Perfusion medium

3. Surgical procedure for liver perfusion

4. Sample preparation

2.4 ME TECHNIQUES USED IN INTACT ANIMAL EXPERIMENTS

1. Measurement of net glycogen accumulation in vivo.

' .2. Measurement of the enzymes of glycogen metabolism in response to hormones.

2.5 ANALYTICAL METHODS

1. Glucose and glucose polymer determination.

2. Carbohydrate metabolite determination.

3. Determination of nitrogenous compounds.

Assay for the enzymes glycogen synthetase and phosphorylase.

48

2.1 THE PREPARATION OF ANIMALS

Albino, male, Sprague -Dawley rats of CFY strain derived

from Carworth, Europe were bred in the Biochemistry Department of

Imperial College; the male stock were renewed every three months,

ensuring a close genetic relationship to the foundation stock. Animals

weighing between 170 and 220g were allowed free access to a standard

(Thompson?s) cereal diet, and water. They were maintained on a light/

dark cycle of 12h (daylight period 06,00h - 18.00h GMT), in a relative

humidity of 55%, and in a temperature of 19-23 °C.

Some rats were starved from 5p.m. until used at 10a.m. or

2p.m. the following day,but the majority of animals were starved for

'48h from 10a.m.

Diabetes were induced by injecting 75mg/kg streptozotocin

(dissolved in about 0.25m1 0.01M citrate - Na pH 4.5) into a tail vein

of a rat under ether anaesthesia. The initial weight (about 210g) on

injection was noted and the animals weighed on subsequent days at a

comparable time-of day (Fig. 2). Four days after injection the blood

glucose of the animal was determined by sampling from a tail vein, and

only if an elevated blood glucose (>12mM) and loss in body weight was

observed, was the rat used in subsequent experiments. It should be

noted that if the animals were not checked at the same time of day each

day, a false impression of changes could be obtained (Table 1). A

similar weight loss between the morning and afternoon weighings was

observed in both the streptozotocin and citrate injected animals but

there was a significant change in the blood glacose of the diabetic

animal compared with the normal.

1

2

4 DAYS

Fig. 2 Growth curve of streptozotocin and control injected

rats

Streptozotocin (75mg/kg) was injected in citrate buffer 0.0IM p114.5

into a tail vein on day zero (0) or citrate buffer alone was injected (0).

Rats were weighed at a comparable time each subsequent day. Results are

mean values S.E.M. of 8 streptozotocin injected animals and 3 controls.

Table 1. The effect of time of day on rat body weight

and blood glucose concentration.

Streptozotocin (75mekg), in citrate buffer 0.0IM pH4.5., was injected

intravenously, or citrate buffer alone. After 4 days the rats were

weighed and the blood glucose determined at the times shown. Each result

is from one observation.

11.30 a.m. 2.30 p.m.

Streptozotocin

Rat No.

1

2

3

1

2

vb. (g)

185

195

212

236

236

Blood glucose 01

20.4

21.6

21.6

6.0

5.2

(mM wt. (g)

180

193

200

229

231

Blood glucose

14.8

16.0

19.4

6.4

5.3

injected rats.

Citrate injected rats.

This was to be expected as animals rarely eat during the day, and

starving is known to partially overcome the elevated blood glucose in

the diabetic state. The rats were starved in separate cages for 48h

prior to use. The rats were not insulin-maintained and some deaths

did occur, especially during starvation. All animals were used within

seven to eleven days of the injection.

Bilateral adrenalectomy was performed' under ether anaethesia

by means of'two lateral incisions. If both adrenal glands were not

clearly removed, in their entirety, the animal was discarded. After

the operation the rats were kept on 0.9% (W/V) sodium.chloride and

weighed every dayI at the same time of day. Their initial weight was

about 165g. (Fig. 3)$ and adrenalectomised rats gained less weight than

controls. Some seasonal variation in the growth of adrenalectomised

animals was noted (Table 2), the animals gaining more weight in the

winter than in the summer. The animals were starved in separate cages

for 48h prior to use, during which some fatalities did occur. All

animals were used within seven to eleven days of the operation.

205

195,

0 *--‘185

I-

W175

165

2 155

2

DAYS

1 3

Fig. 3 Growth curve of adrenalectomised, sham-operated, normal

on 0.9% Nacl and normal rats.

Bilateral adrenalectomy was performed as described in the text and the

rats weighed at the comparable time of day, each subsequent day. Results

are mean values + S.E.M. of at least 24 adrenalectomised animals (0), 8

sham-operated rats maintained on 0.9% Nacl (0), 3 normal maintained on

Nacl (C) and 3 normal animals W. The first two groups of experiments

were carried out in the summer and the last two in the winter.

53

Table 2. Seasonal variation in the growth of

adrenalectomised rats.

Bilateral adrenalectomy was performed as described in the text,

and the rats weighed at the same time of day each day. Results are

means.+ S.E.M.s

Month

with the number of observations in parentheses.

Weight on Weight after Chan e in operation CO 4 days (g) weldL_ )

May 171+2 1641, - 7 (4)

June 164+2 16219 - 2 (4)

July 162+2 — 164+2 + 2 (5)

September 163+2 162+2 1 (18)

October 174+2. 182±3 8 (12)

November 167+2 1682:3 (5)

January 182+2 186+2 4 (12)

February 1.610 173+4 + 12 (5)

March 165+1 175+2 + 10 (6)

54

202 SOURCES OF MATERIALS

All reagents were Analar grade and obtained from Hopkins and

Williams (Ghadwell Heath, Essex), May and Baker (Dagenham, Essex), BDH

(Poole, -Dorset) or Fisons (Loughborough, Leicestershire) unless otherwise

stated. Enzymes for analytical purposes, nicotinamide-adenine

nucleotides, sugar phosphates, pyruvate (mono-sodium salt), UDPG (disodium

salt) and AMP were obtained from C.F. Boehringer Corp Ltd., (London).

Oleic acid, lactic acid, fructose, amino acids, Trizma base and the

hormones hydrocortisone (-21-sodium succinate), vasopressin (8-arginine

and 8- lysine oxytocin and adrenalin (bitartrate) were bought from

Sigma (Kingston-upon-Thames, Surrey).

(8- Arginine)- vasopressin (Grade VI) obtained in solution form

was prepared from synthetic vasopressin of activity about 360 units/Mg;

the concentration of each batch, checked by assay of its antidiuretic

effect in the ethanol loaded rat (carried out by M. Forsling) was about

80% of the stated activity, this factor was taken into account in

calculating the concentration of vasopressin. Synthetic (8 -lysine) -

vasopressin (Grade IV : essentially oxytocin-free) was a powder of

activity 70-100 units/mg and oxytocin (Grade III : synthetic) was in

aqueous solution and reported to be free of vasopressor activity.

Glycogen (rabbit liver) and some amino acids were from BDH, glycerol and

glucose from Fisons. Radiochemicals were obtained from the Radiochemicals

Centre (Amersham, Bucks). Heparin was from Evans Medical Supplies

(Liverpool) and Nembutal from Abbott (Queenboraugh, Kent). Streptozotocin

was prepared and donated by E. Karunanayake (Imperial College, London)

or bought from Upjohns (Michigan, U.S.A.) and recrystallised in ethanol.

55

Insulin was the highest grade commercial ox preparation from Burroughs

Wellcome (Dartford, Kent) and glucagon (crystalline) was obtained from

Eli Lilly (Indianapolis, U.S.A.). Anti-insulin (and control) serum / Dept.

was prepared in the Biochemistry/at Imperial College from guinea-pigs

(Mhnsford, 1967).

56

2.3 THE TECHNIQUE OF LIVER PERFUSION

2.3.1 Perfusion apparatus

The apparatus was essentially that described by Hems et al.,

(1966) based on the designs of Miller et al., (1951) and of Schimassek

(1963). Perfusions were carried out in a thermostatically controlled

cabinet with sash Perspex windows. The contact thermometer in the cabinet

was maintained at 37°C but due to the positioning of the fan, at the top

of the cabinet, the medium entering the liver was a degree lower at

35-36°C. In order to quickly restore the steady temperature after any

____ loss of heat due to the opening of the window for sampling, an

additional fan was placed in the cabinet which could be independently

controlled. The perfusion medium was constantly mixed by magnetic stirrers

built into the floor of the cabinet and small apertures were made in the

sides of the cabinet for the gas supply. This assembly thus provided

a constant temperature environment for perfusion.

The arrangement of the glassware and tubing was as shown (Fig. 4).

'The perfusion medium was pumped from a collecting vessel by a MERE roller

pump (supplied by Watson Marlow Ltd., Cornwall, U.K.) and: passed through

a plastic mesh filter, taken from a disposable blood transfusion set. It

then passed to an oxygenator which was maintained in a vertical position,

facilitating the even flow of perfusion fluid over the surface. The

bulbous nature of the oxygenator increased the surface area of the medium

available for gas exchange. The gas was saturated with water by

bubbling through a wash bottle fitted with a sintered-glas3distributor,

and entered the oxygenator at the bottom and left through an outlet at

the top. At the base of the oxygenator was a small resevoir of perfusate

which was kept at a constant height by an overflow tube leading to the

Filter

✓ Gas

outlet

Oxygenator

Gas in via

wash bottle

Platform for

animal

Outflow to liver

Outflow from liver

Roller pump

Collecting vessel

Fig. 4

aratus for the erfusion of rat liver. • •

57

58

collecting vessel. The height of this resevoir could be adjusted

to give a hydrostatic pressure for optimum flow rates without the liver

swelling. The height was kept at 18 cm. The input to the liver led

from the bottom of the oxygenator via a length of silicon tubing

supplied with a roller clamp, enabling fine adjustments of flow. Th6

perfusion medium from the liver was then returned to the collecting

vessel, and ,the recirculation continued.

59

2.3.2 Perfusion medium

In all experiments the perfusion medium initially consisted

of 50m1 Krebs-Ringer bicarbonate buffer (Krebs & Henseleit, 1932)

and 10m1 15% (N/V) bovine serum albumin. The albumin (Pentex : Fraction

5 from Miles-Seravac Ltd., Berks) was dialysed from three days against

four changes of gassed Krebs-Ringer bicarbonate before use and then

kept frozen at - 20°C. Normally 28-30mM glucose was present in the

medium.

Red blood cells were obtained from a fed donor rat of more

than 600g (one animal, yielding sufficient blood for one perfusion).

The animal was bled from the aorta under ether anaesthesia and the

blood defibrinated on glass beads (Baron & Roberts, 1963) by a rotary

motion of a siliconised flask. After a period of half-an-hour, allowed

for clot contraction, the supernatant was divided into two portions and

washed twice with 20 volumes gassed Krebs-Ringer bicarbonate containing

5mM glucose, except when gluconeogenesis was being studied. A few

experiments were carried out with whole defibrinated blood. The red cells

were then made up to the original blood volume with bicarbonate buffer.

On addition to the perfusate after the start of perfusion, this gave

a haemoglobin of about 4%. In most cases, glucose was added to give an

initial concentration of 28-30MM. Red cell glycolysis was tested in the

absence of a liver, in the presence of 6mM glucose, and was found to

be negligible. The pH of the medium was tested before and during perfusion

and was 7.3 - 7.5. Haemolysis during perfusion was about 1% /h

determined using Drabkin's reagent : 20/41 of sample was added to 4m1

Drabkints (0.05g KO and 0.2g K3Fe (GN)6 / 1) mixed and allowed to stand•

' for 5 min.

60

The resultant coloured solution was read against water at 540nm;

standard 18% haemoglobin gave an optical density of 0.618.

Substrates or hormones were added as an initial dose to the

perfusion medium about 15min after thebeginningof the perfusion and then

infused with a Delta pump (Watson-Marlow Ltd.) at such a rate as to'main-

tain approximately the concentration. The substrate and hormone

concentrations varied from experiment to experiment and will be stated

where necessary. In a number of experiments a mixture of amino acids

was used. This was kept as a stock solution, at concentrations

approximately one hundred times "physiological". "Physiological" levels

were taken as the following, the values being based on the data of

Scharff and Wool (1966) : arginine 0.1mM, lysine 0.4MM, histidine 0.1mM,

pheayialanine 0.1MM, leucine 0.2mM, isoleucine 0.1MM methionine 0.1mM,

valine 0.3mM cysteine 0.1MM, alanine 0.5mM, glycine 0.4MM, glutamic

acid 0.1MM, serine 0.4mM, asparagine 0.2mM, threonine 0.2mM, glutamine

0.6mM, aspartic acid 0.05mM and tryptophan 0.1mM. Tyrosine was added

separately in powder form, giving a "physiological" concentration of 0.2mM

in the perfusate. The stock amino acid mixture was added to give initial

concentrations of one or four times "physiological" (expressed per vol.

whole blood), and then infused at 3ml/h in some experiments.

In some perfusions a supplemented medium was used (essentially

according to John & Miller, 1969), in which were present initially, in

addition to carbohydrate substrates : insulin (500mU), hydrocortisone 21 -

sodium succinate (lmg) and the above amino acid mixture at about four

times the normal concentrations in fed rats.

61

These constituents were maintained by infusion (1.5m1 or 3m1/h

respectively) of separate solutions (in water) containing insulin,

330mU/ml, plus hydrocortisone, 33014-g/ml, or amino acids (see above.)

2.3.3 Surgical procedure for liver perfusion

The operation was carried out essentially as described by

Hems et al., (1966). In early work the animal was anaesthetised with

an intraperitoneal injection of Nembutal (60mg in 0.1m1/100g body

weight) but most operations were carried out under diethyl ether

anaesthesia.The rat was placedon the operating platform and taped

into position. A beaker containing diethyl ether-soaked cotton wool

was kept over the animals head during the operation procedure. A

horizontal cut was made in the abdomen and the intestines deflected to

the animal's left onto saline-soaked tissue. Heparin (0.2m1 = 200

units) was injected into the inferior vela cava and the injection point

covered with a'tissue. The thin strands of connective tissue between

the right lobe of the liver and the vena cava were cut and a loose

ligature of silk (size3/0) placed around the vena cava above the right

renal vein. A double thread was then tied loosely around the base of

the hepatic portal vein and another single ligature put around the vein,

including the hepatic artery. The portal vein was then cannulated with

a No.17 Frankis -Evans needle (trocar and cannula, Luer fitting), the

needle removed and the double thread tied to hold the cannula in place.

A rapid backflow was usually observed which indicated a good and rapid

operation and that the animal was at the correct stage of anaesthesia.

If there was no backflow the cannula was carefully filled with Krebs-

Ringer bicarbonate so as to exclude air bubbles.

62

The thorax was then quickly opened by a transverse incision

above and along the line of the diaphragm and by two deep longitudinal

cuts towards the head. This flap was then removed above the heart.

A loose ligature was placed around the arterior vena cava and a cannula

of Portex tubing (3.00 x 2.00mm / 2.42 x 1.67mm drawn out to a bevelled

point) inserted through the right atrium into the vena cava and down as

far as the diaphragm. The cannula was tied in place.

The preparation was then connected up to the perfusion apparatus

and the first 20m1. medium discarded. During this time the inferior vena

cava ligature and the single one around the hepatic portal vein were

tied. Care was taken that the ties enclosed the camnulae and did not

impede the flow of medium.

After the discard the platform plus animal was placed on the

collecting vessel, the recirculation of medium started and the washed

red cells added. The whole operation took about 10 minutes but the time

from the insertion of the hepatic portal vein cannula to the connection

to the perfusion medium was less than 2 minutes. An indication of the

success of the operation was the uniform olive-green colour obtained

during the washout and the even red-brown colour observed on addition

of the red cells. An extended interruption of the liver circulation

could cause a patchy liver seen either during wash out or when the red

cells were added.

A cage was put over the liver, covered in tissue soaked in

warm Krebs'-Ringer bicarbonate which was itself prevented from drying by a

polythene sheet on top. The flow rate was then adjusted to give a flow

of 16.8 - 18.4m1 /min; difficulty in obtaining this flow rate gave an

indication of lack of success of the operation. The duodenum was

cannulated to allow free flow of bile. There appeared to be no correlation

between the bile produced during perfusion and the rate of glycogen

63

accumulation although it was usually 0.3 - 0.6m1 after 50min of

perfusion. Five minutes after the start of perfusion the gas was changed

from 02 : CO2 (95 : 5) to air : CO

2 (95 : 5). Early experiments were '

carried out with the 02 : • CO2 mixture only.

2.3.4 Sample preparation

Medium samples were removed from the collecting vessel and

mixed with an equal volume of. egperchloricsacid (usually 2ml-i4ml)

for determination after spinning, of glucose or lactate.

Liver samples were removed by looping a thread around the lobe

to be sampled and tying as tightly as possible without shearing through

the liver. The sample would then be taken with the minimum loss of

medium from the cut surface and enabled at least two biopsies to be

taken from each experiment. The liver sample was promptly frozen in

liquid nitrogen and kept at -20°C for eventual determination of glycogen,

enzymes or metabolic intermediates.

64

2.4 THE TECHNIQUES USED IN INTACT ANIMAL EXPEaIMENTS

2.4.1 Measurement of net glycogen apcumulation in vivo

Infusion experiments in intact rats were carried out

as follows. Rats were anaesthetised with an intraperitoneal injection

of Nembutal (0.1m1/100g). Approximately 0.1m1 of 0.9% (W/V) sodium

chloride was injected into a tail vein of the animal; after a few seconds

the syringe was removed. A smallbackflowof blood was observed if

the needle was in the tail vein. The infusion tubing was then connected

to the needle and the pump (Delta : Watson-Marlow, U.K.) was started.

As soon as possible after the infusion was started a liver biopsy for

glycogen determination was taken by making a small transverse incision

to the left of the animals midline and manipulating a lobe of the liver

(left lateral lobe) to the exterior. The biopsy procedure was as for

the perfusion experiments. A small piece of saline soaked tissue was

then placed in the abdomen and the animal kept warm by the proximity

of a lamp. Sixty minutes after the first sample, the second (median

lobe) was taken, and also some blood for glucose determination,

• 65

2.4.2. Measurement of the enzymes of glycogen

metabolism in reponse to hormones

The experimental system was carefully chosen in order to

reduce stress and thus minimise enzyme changes due to stress. Fed

rats were anaesthetised with Nembutal, the abdomen opened and the

rat kept warm by a nearby lamp for 50min. Preliminary experiments

established that steady basal (lowest) hepatic ca' concentrations were

attained 15-20min after opening the abdomen (Kirk and Hems, 1974) and

thus hepatic metabolism was normal by this time. The hormones were

then injected (in 0.25m1 NaC1) into the hepatic portal vein; glucagon,

8 1.0 g; .adrenalin 1.5 x 10 - mol and (8-arginine)-vasopressin, 10 or

100m units. The control injections were 0.9% NaCl. The needle and

syringe were left in position in the blood vessel to eliminate any loss

of blood and thus hormone. The liver was then removed after various

times, rapidly frozen in liquid nitrogen and kept at -20°C until the

enzymes were assayed.

66

2.5 ANALYTICAL METHODS

2.5.1 Glucose and glucose polymer determination

About 0.5g of frozen liver was ground to a powder and boiled

for a few minutes in 10 volumes of aqueous 30% (W/V) KOH. The sample

(which could be stored at this stage) was then boiled for 30 min , and

the glycogen was precipitated with 3 volumes of ethanol (modification

of the method by Good, et al., 1933) and left overnight at 4°C. The

glycogen was then sedimented by centrifugation at 4°C at 10,000 r.p.m

for 15 mins the supernatant discarded and the pellet taken up in

20m1 water, using a motor driven pestle. A suitably sized sample

(usually 0.15m1) containing 0.05 - 0.31-moles of glycogen-glucose was

then hydrolysed to glucose with 30/A1 amyl oglucosidase

in 25MM sodium acetate 1E4.8 (final volume 1m1) for lh at 37°C (Lee

and Whelan, 1966). There was a 98% recovery of glycogen assayed by

this procedure. When 14C-glycogen was counted, glycogen (15-50/Amoles

of glucose) was washed twice in aqueous 70% (V/V) ethanol, hydrolysed

lml of dilute enzyme overnight and dissolved for liquid scintillation

counting in a toltene-based scintillator fluid (see Section 2.5.4)

Glucose in perchloric extracts of perfusion medium,or blood,

or after the hydrolysis of glycogen,was determined with glucose oxidase

(Krebs et al., 1964). A'lml. sample was prepared containing 0.01 - 0.3

rmoles of glucose and to this was added 2.5m1 of enzyme reagent,

composed of 12.5mg glucose oxidase, 4mg peroxidase and 0.5m1

dianisidine in 95% ethanol, per 100m1. glucose buffer (0.5M Na2HPO

4'

0.5M NaH2 PO4 and 0.1M Tris pH 7.3). This was incubated for lh at

37°C and the resultant brown colouration due to oxidation of diarisidine

read at 440nm against a reagent blank.

67

2•5• • Carbohydrate metabolite determinations

Lactate

Lactate was determined with lactate dehydrogenase (Hohorst,

1963). A stock solution containing o.am hydrazine sulphate 2.0M

glycine and 0.02M EDTA (disodium salt) was stored at 4°C until required.

Each day the pH was adjusted to 9.5 and then made up to 2 volumes with

water. The neutralised sample (usually 50/1) was added to a cuvette

containing 1.5m1 of the above buffer, 0.2ml 15mM (1yo) NAD and 1.25m1

water and the extinction read at 340nm against water. Once the optical

density reading was stable 10)A1 lactate dehydrogenase was added and the

reaction allowed to go to completion, as shown by the attainment of a

steady extinction. Any changes due to reagent blanks were subtracted from

E340 and presuming that the molar extinction coefficient of NADH2 was

6.22 at 25°C in a lcm cell the/i -11101 lactate in the cuvette was

calculated. The validity of the assay was checked with occasional use

of"standard lactate solutions.

Glucose 6-phosphate and uridine diphosphoglucose

Glucose 6-phosphate and UDPG were determined as described by

Hohorst (1963) and Mills and Smith (1963) respectively. The analyses

were carried out in sequence in the cuvette.

Liver samples were homogenised in 6% (0) perchloric acid

(1+4 volumes) (Hems & Brosnan, 1970) centrifuged and the pH of the

supernatant adjusted to 7. After standing at 0°C for lh the KC104

precipitate was removed by centrifugation and the supernatant assayed.

(a) Glucose 6-phosphate

lml of the neutralised extract was added to a cuvette

containing 0.9m1 0.IM- Tris -HC1 pH 7.8, 0.0Pm1 0.5M MEC12,

0.02m1 0.25M cysteine (freshly made each day)

and 0.05m1 NAT2. The extinction was read at 340nm against water

and once the reading was stable 2041.41 glucose 6-phosphate dehydrogenase

was added and the final optical density reading taken after completion

of the reaction. The same procedure was applied to tissue and reagent

blanks. Thelt.mol glucose 6-phosphate in the cuvette, and thus

original sample, was calculated, presuming that the molar extinction

coefficient of NA1PH2 was 6.22 at 25°C in a lcm cell. The validity of

the assay was checked by the use of standard glucose 6-phosphate

.solutions; cysteine did not interfere with the assay.

(b) Uridine diphosphoglucose. (.2m3 irn

0.01m1 phosphoglucomutaseiwas added to the cuvette after

completion of the above reaction. No change in optical density reading

at 340nm was observed. 0.02m1 0.I17 potassium pyrophosphate was then

added and no change in E340

noted. Finally 0.02m1 UDPG pyrophosphorylase

was added to the cuvette and the enzymatic reaction allowed to go to

• completion. The same procedure was applied to reagent and tissue blanks;

the recovery of UDPG was 85%. The UDPG in the cuvette and liver sample

was calculated from the molar extinction coefficient of NADPH2 of 6.22

at 340nm.

69

205.3 Determination of nitrogenous compounds

Amino acids.

Perfusion medium was deproteinised with 20 sulphosalicyclic

acid, centrifuged at 3,000r.p.m for 5 min , and the supernatant diluted

and assayed for amino acids.

The amino acids were analysed by means of automated ion

exchange chromatography (Thomas, 1969), the separation and quantification

of each amino acid being completely automatic. The sample was loaded

onto a cation exchange column (Zeocarb 225) from the sampler via a

chromatography pump. This gave a defined flow rate and column pressure.

After electronically programmed elution of amino acids from the column with

an acid to base buffer gradient, fractions of the eluent, separated into

discrete portions by regularly placed gas bubbles, were mixed with nin-

hydrin. The intensity of the reaction was measured colorimetrically,

monitored by means of a logarithmic recorder and quantitated by means

of an integrator. A standard set of amino acids was run with each batch

of samples and the amounts in each sample calculated on the basis of peak

area.

Urea.

The method used for the determination of urea was based on the

principle that urea reacts directly with diacetyl monoxime under

strongly acidic corwi.tions to give a yellow condensation product.

The reaction can be intensified by the presence of ferric ions and

thiosemicarbazide and will then occur without such concentrated acid

reagents; the red coloured complex so formed is more linear with

concentration (Marsh et al., 1965).

Cell free perfusion medium samples were assayed for urea using

the above colour reaction incorporated into an autoanalyser system.

2.5.4 Assay for enzymes glycogen synthetase

and phosphorylase

These enzymes were determined essentially as described by

Das and Hems (1974).

Glycogen synthetase

This enzymes exists in two forms : "a" and "b", usually

assayed as "a" and "a" plus "b". 200-300mg powdered tissue were

homogenised in a Vortex mixer with 6 volumes of cold buffer containing

50MM Tris-HC1, 10mM EDTA and 50mM KF pH 7.5.. Two methods were

employed for the synthetase "a" assay : (i) Method 1 : the extract

was spun at 9,500 r.p.m at 4°C for 15min and the supernatant used for

assay. The assay mixture contained 6.67mM U1PG, 10mg/m1 glycogen,

50mM Tris-HC1 and 5mM EDTA i 7.8 (final concentrations 2/3 those stated);

(ii) Method 2 : the extract was not. spun and the UDPG was increased

to 18mM and 1501 sodium sulphate was added (final concentrations 2/3 those

stated). The mixture for the total glycogen synthetase ("a" plus "b")

assay was as for "a" (Method 1 or 2) but 7.201 glucose 6-phosphate pH

8.0 (final concentration) was present and no sodium sulphate was added.

14C-labelled MPG was added to each mixture to give 40,000 - 80,000 d.p.m

10.1m1 mixture. 0.1m1 of each assay mixture was incubated with 0.05m1

of enzyme at 30°C for 15 min (Method 1) or 10 min (Method 2), the

reaction was stopped by the addition of lml ice-cold 6% (V/V) TCA

containing lmg/m1 glycogen and 2m ml LiBr. Glycogen was precipitated

by 2 volumes of 95% (V/V) ethanol and left overnight at 4°C. The

sample was filtered on a glass filter (Millipore). The assay tube

washed 4 times with 3m1 66% ethanol in water (V/V) and the paper twice

with 2m1 The filter with precipitate was put in a scintillation vial

• and lml of 0.1M sodium acetate pH 4.8 containing 5)&1 amyloglucosidase

added and the vial was left overnight.

71

16m1 scintillation fluid (2.51 2-methoxyethanol, 2.51 toluene, 15g

butyl PBD and 125g naphthalene) were added and the vials counted in

a Packard liquid scintillation spectrometer. Synthetase rates were

calculated as mol glucose transferred from the specific radioactivity

of the precursor.

Glycogen phosphorylase

50-100mg of powdered tissue were homogenised in a Vortex

mixer with 20 volumes of cold buffer containing 35mM 0:- glycerophosphate,

30mM cysteine, 1mM EDTA and 20MM NaP ; final pH was 6.1. This buffer

was made up each day. In some determinations the extract was spun at

9,500 r.p.m at 4°C for 15min, and the supernatant used for assay. The

assay mixture contained 32mM glucose 1-phosphate, 2% glycogen, 2mM AMP

(final concentrations half those stated) and 14C-labelled glucose

1-phosphate to give 40,000 d.p.m/0.2m1 mixture. AMP was included in

the assay to systematise the contribution of phosphorylase wb" to the

activity recorded.

0.2m1 of the assay mixture was incubated with 0.2m1 of enzyme

extract at 3000-for 10min. The remainder of"the procedure was as for

glycogen synthetase.

CHAPTER THREE

RESULTS

73

3.1' THE CHARACTERISTICS AND CONTROL OF HEPATIC GLYCOGEN

SYNTHESIS IN THE NORMAL 48H- STARVED RAT

1. The validation of sequential liver samplin

in the perfusion.

2. The role of. glucose and gluconeogenic precursors

in glycogen deposition in the perfused liver.

The role of insulin and fatty acids in hepatic

glycogen metabolism.

4. Characteristics of glycogen synthetase and

phosphorylase in the liver.

Control of glycogen synthesis in the perfused

liver of normal starved rats.

6. Hepatic glycogen accumulation in the intact rat.

74

3.1 THE CHARACTERISTICS AND CONTROL OF HEPATIC GLYCOGEN

SYNTHESIS IN THE NORMAL 48H- STARVED RAT.

In order to study the control of glycogen synthesis in the

perfused liver, a pre-requisite was the achievement of rates of net

glycogen synthesis, at rates similar to those observed in vivo. Such

rates had not previously been attained in perfusion experiments (e.g.,

see review by Exton et al., 1970). To facilitate this objective,

net glycogen synthesis was measured in perfused livers from starved

rats, so th6A proportional changes in glycogen would be greater (as the

initial glycogen content of livers is lower after starvation). In

this section, the characteristics and control of net glycogen accumulation

in the perfused liver are described.

3.1.1 The validation of sequential liver sampling

in the perfusion

In order to measure glycogen synthesis in the liver by a

sequential sampling procedure it was necessary that the glycogen contents

of different lobes of the liver could be validly related to each other,

i.e., that measurements in glycogen content in one area of the liver should

resemble those in other areas. Biopsies could not be removed in sequence

from within a single lobe, for technical reasons, and also because once

the first liver biopsy had been taken, the remaining part of that lobe

would have an interrupted blood circulation due to the ligature, and would

not represent what may have occurred had it been left intact.

When the major areas of the liver were sampled simultaneously

in intact rats the glycogen contents (5-601xmol of glucose g of fresh

liver) were usually within 10-20% of each other. This was also true

after a short period of perfusion (Table 3) but differences between

• Table 3. Glycogen content and synthesis in the major lobes of perfused liver from 48h- starved rats.

Livers were perfused with bicarbonate -albumin- saline plus fresh defibrinated whole rat blood, containing

glucose (30mM). After 15min, a single dose of pyruvate, serine and glycerol was added (each initially 5mM). Other

details are given in the text. Results are means + S.E.M. with the number of observations in parenthesis.

Glycogen content (tkmol glucose/g fresh liver)

Lobes sampled simult- Lobes sampled sequentially aneously after 20min during perfusion. perfusion.

Rate of glycogen synthesis calculated from sequential liver samples.

(p.mol glucose/min per g)

20 min. 50 min.

Median lobe : Median lobe : Leftlaterel lobe :

13.2 + 2.0 (3)

22.4 + 4.8 (11) 47.0 + 5.0 (11) 0.82

Left lateral lobe : Median lobe : Caudate lobes :

15.2 + 1.5 (3) 22.4 + 4.8 (11) 39.0 + 5.4 (11) 0.55

Combined naudate lobes : Left lateral lobe : Median lobe :

12.3 ± 3.7 (3) 14.6 3.5 (5) 28.3 ± 4.0 (5) 0.46

76 •

the glycogen content of the lobes was observed after longer perfusion.

The apparent rate of glycogen synthesis was greatest in the left lateral

lobe. Since the initial glycogen content of this lobe could be

reasonably estimated by sampling any other part of the liver, the

standard procedure in subsequent experiments was to sample the median

lobe intially, followed by the left lateral. The apparent differences

in the rates of synthesis between the lobes (Table 3) could be explained

by the handling of lobes during sampling. This would be especially true

when the left lateral was being sampled first, followed by the medians

as the latter lies on top of the former and would have to be deflected

towards the thorax in order to ligature the left lateral lobe. It should

be noted that these perfusions were done with whole defibrinated

rat blood and the rates of glycogen deposition were higher than those

obtained when washed rat red cells were used (cf Tables 3 & 4).

77

3.1.2 The role_of_glucose and gluconeogenic precursors

in glycogen deposition in the perfused liver.

The circulating precursors of hepatic glycogen have not been

fully identified (see Introduction Section 1.3). Since the level of

the enzyme glacokinase is reduced on starvation, glucose may not be, the

major carbon source for hepatic glycogen in the starved animal. Glycogen

accumulation was therefore studied in the presence of glucose and

gluconeogenic substrates and their importance as glycogen precursors

assessed.

When the livers from 48h- starved rats were perfused with 30mM

glucose alone, low rates of glycogen accumulation were obtained (Table

4) and glucose uptake of 0.6tkmol /min per g liver was seen. Higher

rates of glycogen synthesis were observed if gluconeogenic substrates

were added to the perfusion medium in addition to the glucose. A standard

mixture of pyruvate, serine and glycerol (each initially 5mM) was used

due to the known rapid rates of gluconeogenesis obtainable from such

substrate combinations (Ross et al., 1967). This mixture provided a

'substantial quantity of carbon atoms for glycogen synthesis without the

risk of inhibitory concentrations of any one substrate, If this mixture

was added 15min after the start of the perfusion, the rate of glycogen

deposition was 0.68)A-mol/Min per g in the presence of 30mM glucose. In

these perfusions no glucose uptake was seen (Table 4).

The glucose concentration in the perfusion medium was critical

for glycogen synthesis. When the concentration of glucose was varied

between 10mM and 40MM maximal rates of glycogen accumulation were seen '

only when the glucose. was 25-30mM (Fig. 5; Table 4). It can be seen

(Table 5) that by increasing the concentration of glucose added to the

medium, the increase in glucose concentration during perfusion became

Table 4. Glycogen synthesis and changes in medium glucose during perfusion of livers from starved rats

Livers were perfused with bicarbonate-albumin-saline containing washed rat erythrocytes and

glucose at various concentrations (initially 10-40MM). When present, a single dose of gluconeogenic substrates

(pyruvate, serine and glycerol; each 5mM) was added after 15min. Glucose in the medium was detected in

duplicate or triplicate. Other details are given in the text. Results are mean values + S.E.M. with the number

of observations in parenthesis.

Medium Glucose - (mM) Glycogen Content . • (pmol glucose/g fresh liver)

MIN: 20 35 50 20 50 (medium lobe) (left lateral

lobe).

Approx. initial Gluconeogenic concn. of glucose Substrates in medium

(Elm)

Rate of glycogen Synthesis

(1nmol glucose/ min per g)

10 11.0+1.1 13.3+1.1 13.7+0.6 7.1 8.3 0.04+0.02 (3)

20 18.1+1.0 19.3+1.0 20.5+0.9 12.1 18.1 0.20+0.06 (7)

30 27.9+0.6 28.9+1.9 28.5+0.6 21.5 42.1 0.68+0.05 (11)

40 38.2+0.8 41.6+1.0 42.8+1.1 13.6 21.5 0.27+0.05 (6)

30 31.6+1.8 30.2+0.1 28.7+1.7 29.8 -- 35.1 0.17+0.09 (5)

ou

a a a

a

79

a

Q0

ra

a

a

a

a

a a a

a

20 MEDIUM GLUCOSE (mM)

40

Fig. 5 Rates of glycogen synthesis at various glucose concentrations

in the perfused liver of starved rats.

Livers of starved rats were perfused with the standard medium,

containing 040mM glucose. After l5min,gluconeogenic substrates were

added (see Table 4). Glucose concentration was measured after 20min of

perfusion, and glycogen synthesis between 20 and 50min. Each point

represents a single perfusion.

Table 5. Calculated total synthesis of glucose during perfusion of liver from starved rats

The amounts of glucose and liver glycogen formed have been calculated from the results in Table 4 and Fig, 5.

Metabolic changes between. 20 and 50min perfusion

Approx. initial Gluconeogenic Change in Medium Glucose Release Glycogen Synthesis Total Glucose con= of glucose in Substrates Glucose (mM) (y.mol/g) ol of glucose/g Synthesised medium (mM) f liver) /30min per g

0 + 3.5 22.4 0 22.4

10 + 2.7 20.6 1.2 21.8

20 2.4 18.0 6.0 24.0

30 + 0.6 5.6 20.4 26.0

40 + 4.6 35.3 8.1 43.4

30 - -2.9 -23.1 5.1 -18.0

81

less extensive, and an increase in glycogen-glucose occurred. However,

the total glucose synthesised did not appreciably alter, and so it would

appear that the glucose was directing the products of gluconeogenesis to

glycogen rather than to glucose. In other experiments with 40thM glucose

plus gluconeogenic precursors (Table 5; Fig. 5) the total glucose

synthesised was greater than that at lower glucose concentrations, and

yet glycogen synthesis was submaximal. These experiments showed that

glucose in the perfusate did not inhibit gluconeogenesis (in the sense

of total glucose synthesis).

The time course of glycogen deposition was investigated under optimal

conditions with the standard liver sampling technique, i.e., median

followed by left lateral lobe. In one group of perfusions the samples

were taken at 5min and 35min, instead of the normal 20min and 50min.

The rate of glycogen synthesis during the first 35min was not as rapid

as that seen between 20 and 50min (Fig. 6),and was possibly due to the

adjustment of liver function after the operative procedure, and not a

true lag in glycogen synthesis.

Since there was no' net uptake of glucose by the liver when

gluconeogenic precursors were added to the perfusion, the net source

of carbon atoms for the accumulated glycogen must have been the

gluconeogenic substrates. Ratesof gluconeogenesis of 1.4 pmol/min per g

were observed in the absence of added glucose (Fig. 7) which showed. that

gluconeogenesis was sufficient to support glycogen deposition, and that

the total glucose synthesised in 30min was of the same order as that

during glycogen accumulation. The rate.fell off with time, probably due

to the decline in the concentration of the substrates.

When glycogen deposition occurred in optimal conditions, i.e.,

in the presence of gluconeogenic precursors and 30mM glucose, there could

have been formation of free glucose by gluconeogenesis and then uptake of

glucose to form' glycogen, in which case there would be no significant net

alteration in medium glucose. The extentof this process

50

40

82

10 20 30 40 50 TIME (MIN)

Fig. 6 Time course of glycogen synthesis in the perfused liver.

Livers of starved rats were perfused as described in Table 4;

the initial glucose concentration was 30mM. Liver samples were taken

at 20 and 50min (E3; 11 perfusions; gluconeogenic precursors added after

15min) or at 5 and 35min (o; 3 perfusions; gluconeogenic precursors added

after lmin). The bars represent the S.M.

83

10 20 30 40 50 TIME (MIN)

Fig. 7 Time course of glucose formation in the perfused liver.

Livers of starved rats were perfused as described in Table 4, except

that no glucose was added. Gluconeogenic substrates were added after 15min.

The larger part of the median lobe was removed after 20min , as in the

standard procedure for measuring glycogen synthesis; its glycogen content

was less than 5pmol of glucose/g. Results are the mean values ± S.E.M.

of 3 perfusions of livers whose average weight (after liver biopsy at 20min) was 5.5g.

84

was igyestigated using (U-14C) glucose. The incorporation of 14C

into liver glycogen (Table 6) showed that assuming that the specific

radioactivity of the medium glucose did not change appreciably, about

one-third of the total glycogen synthesised was derived from glucose

(which was replaced by gluconeogenesis since its concentration did not

fall).

Glycogen accumulation, at rates which occur in vivo, had

thus been observed in the perfusion when the medium contained 30mM

glucose and glycerol, serine and pyruvate (each 5mM).These precursors

were added to the perfusion with glucose in different combinations and

concentrations to see whether any one precursor was the major substrate

for glycogen formation. It would appear that no single substrate was suff-

icient to support significant glycogenesis (Table 7a). The addition of •

10mM substrate to the perfusion medium produced higher rates of synthesis

than when 5mM were used, except in the case of glycerol when the higher

concentration inhibited.Mhen two substrates were added to the perfusion

medium (with 30mM glucose) the rates of glycogen accumulation were

faster than in the presence of each substrate separately (Table 7b) and

the rates appeared to be additive. This also seemed to be the case when

three or more percuxsors were added to the perfusion (Table 7b). No

single substrate thus supported significant hepatic glycogen accumulation,

perhaps with the exception of 5mM glycerol or 10mM serine, alanine or

pyruvate. It seems likely that the net rate of synthesis obtained with

glycerol, serine, pyruvate and glucose,was due to the sum of rates from

each component separately.

85

Table 6.

Incorporation of 140 from (U-140) glucose (30mM) into

glycogen of perfused liver from starved rats.

Livers were perfused as described in Table 4, (30mM glacosb

and gluconeogenic substrates); (U-140) glucose was added to the medium

at 15min. Results are the mean values of three perfusions, in which

the average initial specific radioactivity of the glucose in the medium

was 6.36 x 103 d.p.m/pmol.

Medium 140-glucose (d..p.p/m1) 193,300

Glucose (d.p.m4amol) 6,360

Glycogen in liver l/g) 20min liver sample 37.0+12.0 50min liver sample 59.1+15.5

Net glycogen synthesis (pmol/g/min)

0.74

d.p.m/pmol glycogen 20min liver sample 62.3

50min liver sample 644.0

pmol 140-glucose incorporated into glycogen/g liver. 20min liver sample

003

50min liver sample 5.4

Rate of 140-glucose incorporated into liver glycogen ol/g/min)

0.17

Table 7a. Rates of net glycogen accumulation in perfusions with single substrates

Livers were perfused as described in the text. 30mM glucose was present in the medium and

substrates were added at 15min: any further additions were made at 25 and 35min. Continuous infusion was at

3-4ml/h. Results are means + S.E.M. with the number of observations in parenthesis.

Type of addition (mM) Rate of glycogen -accumulation (Jumol of glucose / g / min)

Glycerol Lactate Pyruvate Serine Alanine

Initial,5 0.42+0.12(5) - 0.09(2) 0.20+0.10(3)

Three additions, 5 0.48(2) 0.42(2) _

Initial 5, plus 0.15+0.06(5) 0.12+0.09(3) 0.02(1) infusion (conc. indicated)

(0.IM) (0.5M) (o.am)

Initial 5, plus two additions, 2.5

0.27+0.10(4) 0.43+0.08(4) 0.30+0.01(3) 0.49+0.16(5) 0.43+0.07(3)

Initial 10, plus infusion (1.0M)

0.23+0.15(3) 0.56(2)

Table 7b. Rates of net glycogen accumulation in -perfusions with substrate combinations

Details are as in Table 7a.

Group Substrate present (initial mM) Rate of net glycogen accumulation ol of glucose/g/min) Glycerol Lactate Pyruvate Serine

1. - - 5 5 0.32+0.32 . (3)

2. 5 - 5 - 0.43+0.08 (4)

5 5 0.65+0.05 (4) (infusion 0.511) (infusion 0.511)

4. 5 0.55+0.09 (4)

5 5 0.68+0.0 (11)

6. 5 5 5 0.52 (2)

88

3.1.3 The role of insulin and fatty acids in

hepatic glycogen metabolism

Due to the metabolic interest of fatty acids and insulin,

their influence on glycogen synthesis was investigated. In optimal

conditions sodium oleate (1mM initially) did not significantly affect

glycogen accumulation(Table 8) although there was a small decrease in

the rate of synthesis. Insulin also had no effect under these

conditions, although in suboptimal conditions (absence of gluconeogenic

precursors, or if glucose was less than 30mM), there was a moderate

stimulation of glycogen deposition ( cf Tables 8 & 4). This apparent

stimulation could have been due to the inhibition of glycogenolysis by

insulin, rather than an actual increase in the rate of synthesis.

Table 8. Influence of added sodium oleate and insulin on glycogen synthesis in the perfused

liver from starved rats.

lavers were perfused as described in Table 4. When present sodium oleate,was initially

1mM (after 15min of perfusion). Insulin was added at 10min and subsequently at 10min intervals, as 0.1m1 of a

1U/M1 solution. Other details are given in the text. Results are mean values + S.E.M., with the number of

measurements given in parentheses when these are different from the number of perfusions.

Gluconeogenic Other No. of Approx.. Glucose in medium (mM) substrates additions perfusions initial

conc. of glucose () 20min 35min 50min

Glycogen content Oamol of glucose /g fresh liver)

After 50mIn (left lateral lobe)

Rate of glycogen synthesis Oamol of glucose/Min per g of fresh liver) After

20min (median lobe)

3 20 22.2+0.6 21.4(2) 21.8+0.7 11.0 21.3 0.35+0.11

Insulin 5 20 19.1+0.9 19.0+0.9 18.7+1.0 22.7 31.3 0.28+0.11

Insulin 5 20 19.4+0.4 21.3+0.4(3) 21.6+0.7 22.3 39.7 0.58+0.14

Insulin 6 30 27.9+0.9 27.5+1.0 27.5+0.8 39.6 51.6 0.40+0.08

9 30 28.4+0.3(6) 30.1+0.5(4) 30.4+0.5 23.6 43.3 0.66+0.05

Oleate 5 30 29.0+0.4 30.6+0.3 30.2+0.5 26.6 43.0 0.54+0.05

Insulin' 5 30 26.1+0.7 27.4+0.4(4) 27.5+0.6 37.6 58.5 0.69+0.09

Insulin, oleate

3 30 30.2+1.6 29.6(2) 30.8(2) 14.7 29.7 0.50+0.10

0,

90

3.1.4 Characteristics of glycogen synthetase and

phosphorylase in the liver

As a pre-requisite to a study of the hormonal control

of the regulatory enzymes of glycogen metabolism, properties of these

' enzymes in homogenates were assessed.For this study, radiometric

assays of synthetase and phosphorylase were employed, as described in

the Methods chapter (Section 2.5.4).

The time courses of the glycogen synthetase

and phosphorylase assays

The time course of the enzyme assays was studied using

centrifuged and crude liver (fed rat) homagnates. The glycogen synthetase

assay mixture was as in Method 1. The centrifuged homogenates brought

about 14C incorporation into glycogen in a linear fashion (Fig. 8).

When the crude liver homogenate was assayed, the rate of reaction was

faster initially.

On calculation of the activity of the enzymes in txmol/min

per g, for each assay time, there appeared to be little difference in

% "a" form glycogen synthetase between 5, 10 and 15min although a fall

in activity of this form and total enzyme was observed (Table 9). The

% "a" form glycogen synthetase in the crude homogenate was however,

higher after 5min incubation than at 10 or 15min. There was a noticeable

fall in the activity of phosphorylase with time measured in the crude

liver homogenate (Table 9). A small decrease was also seen in the

centrifuged homogenate.

Due to the more consistent results, (albeit lower activity);

in the spun homogenate, the majority of subsequent assays were carried

out using this enzyme preparation.

91

For Fig..8 see over-leaf

15 5

10

ASSAY TIME MIN

I I

0 5 10 15 0 6 10

7000

A

PHOSPHORYLASE SYNTHETASE "a"

TOTAL

2400 SYNTHETASE

1600

800

1-:=1

n 5000

O

m 0

2 O

3000

0 0 m

1000

Table 9. The effect of different assay times on the activities of hepatic glycogen synthetase and

phosphorylase.

Assay details are in the text. Results are from a single fed rat liver, except for the glycogen

synthetase measured at 5 and 15min which are means of 2 animals. Assays were done in duplicate at 30oC.

Glycogen synthetase activity -method 1) pimol/min per g)

Assay time

Iran

5min. Total OA Half

lOmin. Total % "a"

Crude homogenate Centrifuged honte

0.31 0.10

0.69 0.21

45 48

0.26 0.11

0.81 0.21

33 50

"a"

0.18 0.08

15min. 15min. Total 0.53 0.16

Yo "a"

34 48

Glycogen

3min. 14.70

5.75 phosphor-lase activity

6min. 13.73

5.27

per g)

10min. 11.68

5.21

94

The effect of centrifugation on enzyme activity

Glycogen synthetase was assayed using the mixture defined as

Method 1 and incubation was for 15min for glycogen synthetase and 10min for

phosphorylase at 30°C, or 2min (both enzymes) at 37°C. When the assay was

carried out at 30°C, centrifugation or the homogenate caused loss of activity

of both glycogen synthetase and phosphorylase (Tables 9 and 10). The total

glycogen synthetase of fed rat liver however, decreased more, resulting in

an apparent increase in % "a" form glycogen synthetase. A similar fall in

activity was observed when the assay was done at 37°C (Table l0),although

there was not such a large difference between the spun and crude homogenate

(compared to that at 30°C) when the glycogen synthetase of the fed rat liver

was assayed.

It appears therefore that centrifugation of the liver (from a

.starved rat) homogenate halves the activity of glycogen synthetase and

phosphorylase when assayed at 30°C and 37°C. In the fed liver homogenate

the situation is not as clear, as spinning caused a greater loss of glycogen

activity when assayed at 30°C than at 37°C. This apparent loss of activity

due to centrifugation has been reported especially with respect to

phosphorylase,which sediments with the microsomal fraction in the fed state

but not in the starved (Maddaiah & Madsen, 1968; Tata, 1964). No such

correlation has been observed with glycogen synthetase, although a large

proportion of activity is associated with the microsomal fraction in the fed

state (Maddaiah & Madsen, 1968).

Table 10. The effect of variation in tem•erause and •unification on the activities of he•atic

glycogen synthetase and phosphorylase in the fed and starved rat

Assay details are in the text. The assay mixture for glycogen synthetase was that defined as

Method 1, and incubation was for 15min for glycogen synthetase and 10min for phosphorylase at 30°C, or for 2min

(both enzymes) at 37°C. Results are means of 3 rats + S.E.M.; mean percentages of synthetase "a" were

calculated from % "a" values in individual samples. Assays were done in duplicate.

Fed rat

48h- starved rat

Assay Temperature

30°C

37°C

Liver homogenate

Crude homogenate

Spun homogenate

Crude homogenate

Spun homogenate

Glycogen synthetase (12 mol/min per g)

Glycogen phosPhorylase. Ou mol/min per g)

14.61

+1.47

5.20

+0.93

25.73

+2.24

7.57 +0.91

Glycogen synthetase

mol/min per g)

Glycogen '.phosphorylase ()a mol/min per g)

9.20

+0.95

5.06

±0.94

16.42

+0.90

8.73

+1.04

"a"

0.16

+0.02

0.09

+0.01

o.56

+0.10

0.50

±0.04

Total

0.50

+0.04

0.15

+0.02

1.64

+0.13

1.05

+0.24

% "a"

32

±2

49 ±1

36

+7

51

i)

"a"

0.18

+0.04

0.08

+0.01

0.85

+0.09

0.47

+0.04

Total

0.39

+0.07

0.19

±0.02

1.35

+0.16

0.83

+0.19,

% "a"

45 ±3

43 ±1

63

±1

63

+13

96

The effect of assay temperature on enzyme activity

When crude liver homogenates were assayed at 37°C a 3-4 fold

increase in activity was observed (Table 10) for glycogen synthetase

and a 2 fold increase for phosphorylase,compared to values at 30°C.

The glycogen synthetase activity of the spun homogenate at 37°C was

increased 6-7 fold, and phosphorylase 2 fold.The assay temperature ,the

therefore had a greater effect on/spun homogenate than on the crude

homogenate. This response could imply that the crude extract activity

approximates the ttruel activity of the enzyme and thus increasing the

assay temperature has a less dramatic effect.

97

3.1.5 Control of glycogen synthesis in the perfused

liver of normal starved rats.

In order to assess the role of hormones in the control of

the regulatory enzymes of glycogen metabolism, the properties of these

enzymes were studied in the perfusion. Glycogen synthetase and

phosphorylase were measured in the perfused liver of normal starved rats

under optimal and sub-optimal glycogen accumulation conditions and any

correlation evaluated.

In these conditions, the hepatic concentrations of UDPG and

glucose 6-phosphate were measured (Table 11); the latter value was of

the same order as previously reported for perfused liver of starved

rats (Ross et al., 1967). When maximal rates of glycogenesis were

observed, the concentration of MPG was decreased, compared to that when

lower rates of glycogen accumulation were found (Table 11), suggesting

that the glycogen synthetase reaction was accelerated. This inference

was confirmed, since under these conditions an increased proportion

(60-80) of glycogen synthetase was present in the "a" form (an active

form in vivo). This was true despite the lack of a pattern in total

synthetase activity or actual activity of the "a" form, and suggests

that the % of synthetase in the "a" form may relect its capacity to

bring about net glycogen deposition.There was decreased activity of

glycogen phosphorylase in perfusions with glucose (Table 11) but this

was not sufficient in itself for high rates of glycogen accumulation

to be obtained, unlike the situation when C3-substrates or fructose

were added to perfusate in addition to the glucose. In these latter

perfusions glycogen phosphorylase was decreased in parallel with the

increase in % glycogen synthetase "a".

Additions to perfusion medium

No. of Enzyme assays (amolAin/g of perfusions Glycogen synthetase

(Method 1)

Han Total

% Hatt

None 3

0.14 + 0.01 0.38 + 0.04 38 + 3

Glucose 3 0.18 + 0.03 0.37 + 0.05 48 + 3

03-substrates 4

0.17 + 0.02 0.42 + 0.11 46 + 8

Glucose plus 6

0.20 + 0.01 0.34 + 0.02 61 + 3 03-substrates

Glucose plus 0.16 + 0.02 0.21 + 0.03 80 ± 4 fructose

Glucose plus* 4 0.17 + 0.06 0.20 + 0.05 83 + 7 03-substrates

* Supplemented medium, including insulin, hydrocortisone and amino acids.

t Net rate of glaconeogenesis 1.22 + 0.10 (5)

fresh liver) Glycogen phosphorylase

(centrifuged homogenate)

5.6 ± 1.0

3.6 + 0.3

7.3 ± 1.1

3.4 ± 0.5

1.4 + 0.4

2.3 + 0.5

Table 11. Concentrations of enz es and •athw intermediates durin 1 co:-n s thesis in the .erfused

liver of starved rats.

Livers from 48h-starved rats were perfused with bicarbonate-albmin-saline containing washed rat erythrocytes. Substrates where present were added after 15min. Glucose when added was initially 28mM, and the 07- substrates, lactate, glycerol and pyruvate, were initially 5mM, 3.3mM and 1.7mM (respectively) and then ilifused (3m1/h) in a mixture containing 0.5M-sodium lactate, 0.33M-glycerol and 0.17M-sodium pyruvate. Fructose was initially 5mM, and then infused (0.5M, 3m1/11). Perfusion was for 50min except those with glucose alone : 60min. Enzymes, glycogen and metabolites were assayed in frozen samples of the left lateral lobe. An initial sample (median lobe) was removed, after 20min to measure glycogen. Results are means + S.E.M of the no. of observations indicated; mean percentages of synthetase "a" were calculated from % "a" values in individual samples.

•

Metabolite content Rate of net (nmol/g) glycogen accumulation

UDP- Glucose 6- (p010 1 glucose/min/g) .glucose phosphate

28 ± 7(3) 20(3) 0

96 ± 12(4) 48 ± 8(4) 0.21 + 0.08

89 ± 13(5) 51 ± 6(5) 0

52 ± 11(10) 71 ± 11(10 0.75 + 0.10

0.84 + 0.07

0.54 + 0.19

99

3.1.6 Hepatic glycogen accumulation in the intact

rat.

In order to assess the physiological significance of the

properties of hepatic glycogen accumulation observed in the perfusion

(Section 3.1.2), glycogen deposition was measured in the intact

anaesthetised 48h- starved rat, by infusion of substrates via a tail

vein. The loft lateral lobe was removed first, followed by the median lobe,

as this was technically the easier order of sampling. Separate experiments

(not shown) revealed that in vivo (unlike perfusion), either order of

sampling gave the same rate of net synthesis. Unlike the perfusion

studies, glycogen accumulation was obtained when glucose alone was

infused. However, this result is not incompatible with those obtained

by perfusion as gluconeogenic percursors would have been released by

other tissues (e.g., muscle) in the intact rat. The rate of glycogen

deposition was dependent upon the final blood glucose concentration

(Fig. 9) in a similar way to that observed in perfusion experiments. As

might be expected the time lag observed in glycogen synthesis in the

perfused liver was not as pronounced when liver biopsies were measured

at various. times during in vivo infusions (Fig.10).

100

0.8

0 0

0

.E 0.6 E

0 E

a, _c C

0.4 — c o.) 0 t.) a) 0

0

0.2--

0

0 0

1 I I I 20 40

Blood glucose (mM

Fig. 9 Rates of glycogen synthesis in vivo, at various blood

glucose concentrations.

Glucose was infused intravenously, from 5 to 90min

after anaesthesia. Liver samples were taken after 30 and 90min, and

blood after 90min. Other details are given in the text. Each point

represents a single experiment.

. 0

0 0

0 0 0

0 0

0

60

40 w - N Z 0 u - 7

0)

LIJ g- o 0 •

O -6

>L2" X20 0

101

1

0 20 40 60 80 100

TIME(MIN)

Fig. 10 Time course of clycotIen synthesis in viva.

Glucose was infused intravenously into anaesthetised rats.

In three groups of experiments, each with six rats, and each denoted

by a different symbol, two sequential liver samples were taken : 0,

at 5 and 35min after the start of the infusion, or A , after 5 and

65min, or 0 after 30 and 90min. The blood glucose concentration

at the time of the second sample was 23-36mM.

102

302. HEPATIC CARBOHYDRATE AND FAT METABOLISM IN

THE "FED" RAT

1. Glucose metabolism in the perfused liver of fed rats.

2. Glycogen synthesis in the perfused liver of overnight-starved rats.

3. Fatty acid synthesis in the perfused liver of overnight-starved rats.

103

3.2 HEPATIC CARBOHYDRATE AND FAT METABOLISM IN THE "FED"RAT

The role of glucose and gluconeogenic precursors in

hepatic glycogenesis in the 48h- starved rat has been studied (see

previous Section). The contribution of glucose to the net rate of

glycogen accumulation in the perfused liver was small,being zero in

terms of the net rates (as there was no glucose uptake); unidirectional

uptake was about one-third of the total rate. Since the level of the

enzyme glucokinase is reduced on starvation (Salas et al., 1963; Walker

& Rao, 1964), it was of interest to study the role of glucose and

gluconeogenic substrates in fed rats, where glucokinase and glucose

uptake could have a greater role.

3.2.1 Glucose metabolism in the perfused liver of

fed rats.

In fed rats, hepatic glucose uptake was faster than

in 48h- starved rats (Fig. 11). The existence of net gluconeogenesis

was demonstrated in livers from fed rats perfused with a mixture of

glycerol and pyruvate, plus lactate (maintained by infusion); this mixture

brought about net glucose output, in the absence of a decline in glycogen

(Fig. 11). If lactate was replaced by serine, net glucose uptake was

also reversed by this mixture of gluconeogenic precursors. The rate of

net glucose output in the experiments with the lactate - containing

mixture, calculated from Fig. 11 (disregarding the small amount of

net glycogen synthesis) was about 0.5p.mol glucose/Min /g liver, or

about 4jAmol/Min/iiver.

104

40

E

(I) 0 U _I 20

0

500

O

0 300

O

o

20 60 100 TIME (MIN )

Fig. 11 Glucose metabolism in the perfused liver of fed rats

Livers were perfused with glucose (initially 30mM) as described

in the text. The following additions were made, at the times indicated:

mixture of glycerol and pyruvate plus lactate (40min: see Table 11: A )

or serine (30min : see Table 41V ). In the experiments with the lactate

- containing mixture, glycogen in sequential biopsies was measured,

( G3 , broken line -- -). In two groups of perfusions, no additions were

made, in fed (o) and, for comparison, 48h- starved rats (0). Results

are from three perfusions.

105

3.2.2 Glycogen synthesis in the perfused liver

of overnight-starved rats.

A common procedure is to starve animals (or people)

overnight, in an attempt to stabilise the metabolic events of the "fed"

state. This depletes liver glycogen which permits the study of net

glycogen accumulation. Rates of glycogen synthesis were measured in

perfused livers from such rats, in conditions shown to be optimal

for net glycogen deposition in 48h- starved rats, and the role of

glucose and gluconeogenic substrates assessed.

In perfusions with glucose (NUM, no other substrates),

the rate was higher than in 48h- starved rats (Table 12). The rate of

glycogen accumulation was lowest in perfusions with glycerol-containing

uixture, and was unchanged by insulin (Table 12). This apparent

inhibitory action of glycerol was not due to diminution of the rate of

gluconeogenesis, the rate of which was 1.05+0.121(molAtin/g of fresh

liver (mean + S.E.M. of three perfusions with the glycerol mixture

measured in the absence of added glucose) i.e., more than three times

faster than the-rate of glycogen synthesis (0.281A-mol/min/g : Table 12).

In livers from 22h- starved rats, perfused at 14.00h with glucose plus

lactate, glycerol and pyruvate, the rate of glycogen accumulation was

higher than in 18h- starved rats (Table 12).

The above rates of glycogen synthesis, especially

in the presence of the glycerol-containing mixture, suggested that there

could be an inherent impairment in the capacity for net glycogen synthesis

in overnight-starved rats. However, experiments with anaesthetised intact

animals showed that this was not the situation. Net glycogen

accumulation, measured in sequential liVer biopsies in vivo, during glucose

infusions was 0.77+0.12 (4)tx.mol glucose/g/min at 30mM glucose, which is

Table 12. G1 co en synthesis in the perfused liver of overnight-starved rved rats

Livers were perfused as described in the text at 11.00h unless indicated. Substrates where

present were added after 15min. Glucose was initially 28rE and the gluconeogenic precursors, L(lactate), G(glycerol)

and P(pyruvate) were initially 5mM, 3.3mM and 1.7mM (respectively) and the infused (3ml/h) in a mixture containing

0.5M-sodium lactate, 0.33M-glycerol and 0.17M-sodium pyruvate. Mien the L and P mixture was added, lactate was

8mM and pyruvate 2mM and theywere infused (3m1/h) in a mixture containing 0.8M-sodium lactate and 0.2M-sodium

pyruvate. Fructose was initially 5mM and then infused (0.5M, 3m1/0. Insulin was added (250mU) every 15min.

Results are means +

Period of Additions to No. of Glycogen (,pmol of_glucose/g) Perfusate glucose Calculated rate starvation perfusate perfusions 20 min 50 min (mM) of net glycogen (h) 20 min 50 min accumulation

(median (left lateral ' cumol of glucose

lobe) lobe) /g/min)

18 None 8 6.3 17.6 26.3 23.7 0.38 + 0.09 18 L, G, P 6 2.0 10.6 27.1 29.6 0.28 + 0.05

18 L, G, P plus insulin

2 0 7.7 28.6 27.6 0.26

18 L, P 3 6.8 22.0 24.1 25.3 0.51 + 0.03

22* L, G, P 4 2.0 18.7 27.8 29.8 0.56 + 0.02

18 Fructose 3 5.3 27.0 27.7 32.1 0.73 4. 0.10

48 None 5 29.8 35.1 31.6 28.7 0.17 + 0.09

48 L, G, P 12 23.0 44.6 30.3 - 0.72 + 0.05

* perfused at 14.00h L H O

107

similar to the rate (0.6 - 0.7) in 48h- starved rats (see Section 3.1.6).

Also, if overnight-starved rats livers were perfused with glucose plus

fructose, maximal rates of net glycogen accumulation were observed

(Table 12), again showing that there is no inherent impairment in

glycogen synthesis capacity after starvation.

3.2.3 Fatty acid synthesis in the uerfused liver

of overnight-starved rats

The total rate of fatty acid synthesis was measured with

31120 (Salmon et al., 1974) in livers from overnight-starved rats,

perfused with glucose (30MM, plus the mixture of lactate, glycerol and

pyruvate). The rate was 0.5+0.3 (3)14.mol of long-chain fatty acid/g,

ywhich contrasts with rates of 3 - 5in fed rat livers in comparable conditions (Kirk, et al., 1975). These low rates may be explained by the

decline in hepatic glycogen, which serves as a source of acetyl residues,

and also exerts a stimulatory role in lipogenesis (Salmon et al., 1974).

108

3.3 DIE ACTIONS OF THE HORMONES OF THE

POSTERIOR PITUITARY GLAND ON HEPATIC

GLYCOGEN METABOLISM

1. Stimulation of hepatic glycogen breakdown

by (8 -arginine)-vasopressin and oxytocin.

2. The role of vasopressin in hepatic glucose

metabolism.

The stimulation of hepatic gluconeogenesis

by (8-arginine)-vasopressin.

The action of vasopressin and oxytocin on

glycogen synthesis in the perfused liver

and the intact rat.

5. The effect of vasopressin and other

glycogenolytic hormones on hepatic glycogen

synthetase and phosphorylase in vivo.

109

3.3 THE ACTIONS OF THE HORMONES OF THE POSTERIOR

PITUITARY GLAND ON HEPATIC GLYCOGEN METABOLISM

Antidiuretic hormone (vasopressin) is involved in

the processes of water balance and causes reabsorption of water in

the kidney in order to maintain a constant blood volume, compositiOn

and pressure. Glycogen is stored in the liver in association with

large quantities of water (Fenn, 1939) and since vasopressin blood

levels are increased during shock (e.g., haemorrhagic stress) .44-. was

of interest to evaluate the role of vasopressin, in hepatic

carbohydrate metabolism (see Introduction, Section 1.4.3). Also, since the

role of the posterior pituitary gland in metabolic events is not fully

clear, oxytocin action on the liver was studied.

3.3.1 Stimulation of hepatic glycogen breakdown

by (8 -arginine)-vasopressin and oxytocin

It has been reported (see Introduction, Section 1.4.3)

that vasopressin and oxytocin cause hyperglycaemia by stimulating the

breakdown of hepatic glycogen. It was of interest to study this

action in the perfused liver.

During perfusion of livers from fed rats the glucose

concentration in the medium rose initially from 5mM to 8mM and then

remained steady. If (8 -arginine)-vasopressin was then added in a

single dose to the medium there was a marked rapid efflux of glucose,

which fell off after about 40min (Fig. 12). This decline in glucose

efflux was not caused by lack of glycogen, as shown by glycogen

measurements at the end of the perfusions. The extent of the change in

glucose concentration was dose dependent, over the range 50 -609units/m1

of vasopressin (Fig.13), saturation occurring at 6001.kunits/ml.

110

20 —

1 40

80

120 TIME (MIN)

Fig. 12 Influence of va o ressin on the time course of

glucose output in the perfused liver of fed rats.

Livers were perfused as described in the text. The following

additions were made to the perfusion medium after 40min : vasopressin

at the following initial concentrations (p. units/ml) 1,000,0; 130,A ;300,0;

5,0 , or 0.7m1. of an extract of rat neuro-hypophysis 0 . Results

are from single perfusions.

0.5 VASOPRESSIN (mU/mI

1.0

Fig. 13 Dependence of the stimulation of hepatic glucose

output on vaso ressin concentration.

Livers from fed rats were perfused as described in the text.

After 40min, vasopressin was added, at various initial concentrations.

Each point represents the increment in glucose output during the next

40min , determined for each perfusion.

112

The most likely explanation for the rise in glucose concentration

in the medium was that breakdown of liver glycogen was being

stimulated by vasopressin. This was confirmed by measuring changes

in the hepatic content of glycogen during continuous infusion of

(8 -arginine)-vasopressin (Fig. 14). The rate of glycogenolysis was

calculated from the change in glycogen content of the two sequential

liver samples taken in each perfusion, presuming that no. large

differences in glycogen content exist between the major lobes. The

mean decrease in glycogen content in 60min. . was 49-1-14 (5)/A.mol of

glycogen glucose / g in control perfusions and 121±51 (4) in the

presence of vasopressin. The glycogen breakdown due to vasopressin

(about 7014mol of glucose / g) thus amounted to about 409kmol of glucose

(the average liver weight being 5.6g), which was sufficient to account

for the extra glucose (1501.4mol) which appeared in the medium. There

was no major effect of vasopressin on the lactate concentration in the about 40min

medium. The plateau in glucose efflux observed/after a single dose of

hormone (Fig.12) was still observed during (8 -arginine)-vasopressin

infusion, indicating that a single dose of hormone is sufficient for

a response to be observed, or that high concentrations of glucose inhibit

the vasopressin effect.

Apart from the fact that the posterior pituitary gland

secretes oxytocin as well as vasopressin, they are very similar

polypep-bides in structure and so it was of interest to replace vasopressin

by oxytocin in the above described system. A dose of 5000runits / ml

(10 times the saturating vasopressin dose) was added to the perfusion

medium; glucose efflux was negligible, but was observed at higher

concentrations (Fig.15).

LIV

ER

GLY

COG

EN

400

rn

a) 0

0) 200

'46

0 E

0

12

E E10 D

E

L.1.1 8

••••••••

e■I

0 4

tr) 0

2 -

A 4 ------A

A A

I

1

° - - 0 ----CI 0 • - - - - - • 0

0

113

0 20

40 60 80 . 100

TIME ( MIN)

Fig. 14 Effect of vasopressin on glycogen Content of

the nerfused liver.

Livers from fed rats were perfused as described in the text. After

40min vasopressin was added to an initial concentration of 700 punits /ml, and then infused at 200m units/h. An initial liver sample was

removed at 41min. The average liver weight for the remainder of '

the perfusion was 5.6g. Results are from three control perfusions

(open symbols), and four with vasopressin (filled-in symbols) : (0,0)

glucose; (A, A ) lactate; (0, 0 ) glycogen.

114

18

fc' --0- 9--1-1° A 0-1:1 A,4,/ ;

p,,N"- 0 ,/0 ■ --0 0 —0 0

0

0 20 40 60 80 100 TIME (MIN)

Fig. 15 Influence of oxytocin on the time course of

_lucose out ut in the erfused liver of fed rats.

Livers were perfused as described in the text. The

following additions of oxytocin were made to the perfusion medium

after 40min,at initial concentrations of lm units/m1(0);50m units

/ml (0) ; 100 m units/ml ( ); 500 m units/m1(A) . Results are

from single perfusions.

115

The commercially prepared vasopressin was checked

by a number of procedures. The amino acid composition after hydrolysis

in 6.14 liC1 was that expected of vasopressin thus excluding significant

contamination with additional peptide material. The glycogenolytic

effect of the preparation was largely destroyed by incubation in

10mM-thioglycollate (90min, 37°C, vasopressin 100m units Al), or by

combination of freezing and thawing, and standing at room temperature

for 20h. These results were compatible with the glycogenolytic agent

being vasopressin, which is known to be inactivated by these procedures.

An extract of rat neurohypophysis was prepared (in 0.5m1 of 0.1 M-HCl,

homogenised, centrifuged, and the supernatant neutralised) and on

addition to the perfusion caused glucose output (Fig.12) probably due

to vasopressin)since oxytocin (5m unit/m1) has a much less potent effect

on glucose efflux (Fig. 15).

No detectable alteration in the rate of flow of perfusion

mediumwas observed at any vasopressin concentration tested i.e.,

5 -10001kunitshal.

3.3.2 The role of vasopressin in hepatic glucose

metabolism

When fed rat livers were perfUsed with 30mM glucose,

glucose uptake was observed which was reversed by gluconeogenic

precursors (Section 3.2.1). In the presence of high glucose

concentrations (30mM) net glycogen accumulation is maximal and net

breakdown minimal (Glinsmann, et al., 196A and so this state might thus

be expected to influence the action of catabolic hormones. Such a

possibility was investigated in perfusions containing vasopressin

(Fig.16), which had a potent glycogenolytic action at autoregulatory

(9-12mM) glucose concentrations (Section 3.3.1). At 30mM glucose,

glucose output due to the hormone was negligible (Fig 16), although net

116

40

ID 1:3 El

---,... :1., 0 0 DI

E NN. 0 El ■-• w 0 CI U---....„.......0 D (-51 20`—

D (3 ul

I I I I I 1 40 80 120

TIME (MIN

Fig. 16 Effect of vaso ressin on lucose metabolism in

the perfused liver of fed rats.

Livers were perfused with glucose (initially

30mM) as described in the text. Between 40 and 60min 3 additions

of vasopressin, 110/m1, were made (0); control perfusions (❑ ).

Results are means of two perfusions with vasopressin, and three

controls.

0

117

glucose uptake was halted. A similar inhibition by glucose was observed

during infusion of the hormone (see previous section) when a decline in

glucose efflux was observed after 30mi .

3.3.3 The stimulation of hepatic gluconeogenesis by

(8-arginine)-vasopressin.

The observed stimulation of glycogen breakdown by vasopressin

raised the possibility that the hormone may act in a similar way and on

the same processes as•glucagon. The action of vasopressin on

gluconeogenesis was thus investigated.

The hormone was added in five doses at regular intervals due

to its rapid destruction by the liver (Little et al., 1966). The

concentration of (8-arginine)-vasopressin in the perfusion medium, measured

at the end of the perfusions, was not higher than that produced by each single

addition (M.L. Forsling, unpublished work), hence it was reasonable to

consider the hormone concentrations during the perfusion to be equivalent

to that produced by the initial addition. Gluconeogenesis from an infused

mixture of lactate, glycerol and pyruvate was stimulated by vasopressin

over a concentration range 30-150)u units/ml. (Fig. 17).

A similar stimulation (from 1.61+0.10 (6) to 2.46+0.09 (3)

min/g) mes.observed when two doses of the hormone were added after a

control rate of gluconeogenesis had been obtained with the gluconeogenic

mixture (Fig. 18). The effect of vasopressin on gluconeogenesis in the

absence of added substrate was also tested; no major stimulation occurred

(Fig. 18). Oxytocin (1000).Lunits/m1) had no effect on the rate of

gluconeogenesis from the glucogenic mixture.

O cs)

•■■

0 0

0

O

inc r

ease

0.5

0

2.5

GLY

CO

GE N

2.0

118

50 100 150

280

VASOPRESSIN(,ill ml

Fig. 17 Influence of vasopressin on gluconeogenesis or net glycogen accumulation in the perfused liver of starved rats.

Livers were perfused with gluconeogenic precursors (glycerol, lactate and pyruvate) as described in Table 12. Vasopressin was added at 10min intervals, from 10min after the start of the perfusion, to the concentration shown (abscissa). Two groups. of experiments were carried out: (a) 0 measurement of gluconeogenesis; rates were measured between 20 and 60min. Half the median lobe of the liver was removed at 20min (as in the standard procedure when measuring glycogen synthesis) and the glycogen content was negligible. Each point represents a single perfusion; (b) , measurement of net glycogen synthesis; perfusions contained 30mM glucose and gluconeogenic precursors, and results are means of three or four measurements (bars indicate S.E.M.).

119

T0 1 14

12

40 80 120 TIME (MIN)

Effect of va

(endo enous and from added substrate in the •erfused

liirer of starved rats.

Livers were perfused as described in the text and Fig. 17. Gluconeogenic substrates, glycerol, lactate and pyruvate were added at 20min and then infused (0, 0). Half the median lobe of the liver was removed at 20min : the remaining liver being 3.77 and 3.67g respectively.(8-Arginine)-vasopressin (570,u units/ml) was added at 60 and 75min (o). In one group of experiments (A) no substrates were added and vasopressin (1,000ju units/ml) was added at 40 and 70min; in these perfusions no liver was removed and the mean liver weight was 4.73g. Results are means of three perfusions except for the control period with substrates before vasopressin addition(6) and bars indicate S.E.M.

0

Fig. 18

120

3.3.4 The action of vasopressin and oxytocin on glycogen

synthesis in the perfused liver and the intact rat

It has been shown that net rates of glycogen accumulation in

the perfused liver require the presence of gluconeogenic percursors

as well as glucose (see Section 3.1.2). Yet (8 -arginine)-vasopressin

stimulates both gluconeogenesis (in the starved animal : results above)

and glycogenolysis (in the fed animal : results above). It was therefore

of interest to study the action of the hormone on net glycogen

accumulation in the perfused liver from a starved rat.

Glycogen accumulation was inhibited by (8-arginine)-vasopressin

in the perfused liver (Fig. 17) and the dependence of this effect on

the hormone concentration, was similar to that observed when

gluconeogenesis was studied. In these experiments there was no clear-

cut effect of vasopressin on the total glucose synthesised (i.e., the

sum of the change in liver glycogen plus blood glucose). (8 -Lysine)-

vasopressin also inhibited glycogen accumulation (Fig. 19),the dose

required for half-maximal synthesis being 20p units/ml compared to a

dose of 5011 units/M1 for (8-arginine)-vasopressin. This apparent

difference in sensitivity is diminished when the concentrations of the

hormones are considered on a weight basis (Fig. 19), inhibition occurring

over the range 60-300 pg/ml (vasopressin - like activities of the pure

hormones were taken as 400 U/mg and 250 U/mg, respectively; Boissonnas

et al., 1961).

Under the same conditions as described above, oxytocin inhibited

glycogen accumulation (Fig. 19)

(arginine or lysine forms), the

half-maximal about 25 m units/M

but at much higher doses than vasopressin being 400

saturating dosesAbout/M units/M1 and the

. Glycogen synthesis is thus inhibited

121

103 105 HORMONE IN MEDIUM (log j ml

10'

GLY

CO

GEN

A

Fig. 19 Effect oiLLEi me,son and oxytocin on glycogen

synthesis in the perfused livers of starved rats.

Livers were perfused with 30mM glucose and a gluconeogenic mixture of glycerol, lactate and pyruvate as previously described; glycogen synthesis rates were measured between 20 and 50min. (8 -lysine) -vasopressin (10 and oxytocin (0) were added at 10min intervals from 10min after the start of the perfusion, to the concentration shown. For comparison, the dependence of the effect of (8 -arginine) -vasopressin is included : broken line. Results are from individual perfusions except for controls (0), ' 14 observations.

122

over the range 5 - 400 m units/M1 or 10 - 900 neml (pure hormone has

a reported activity of 450 U/mg ; see Boissonnas et al, 1961).

The results with (8-arginine)-vasopressin in the fed and

starved animals indicate that the starved animal maybe more sensitive

to the hormone.

In order to confirm that the above observed actions of (8 -

arginine)-vasopressin occur in vivo the action of the hormone was

investigated in the intact rat. Although the ideal experiment would

have been to study its effect on fed liver glycogen, the glycogen levels

found in fed animals are variable and it is thus difficult to follow

changes in the intact animal. It was therefore, decided to use the 48h-

starved rat . Vasopressin and glucose were infused into the intact

anaesthetised rat and liver biopsies removed. Inhibition of glycogen

synthesis was observed (Table 13), confirming perfusion results.

3.3.5 The effect• of vasopressin and other glycogenolytic

hormones on hepatic .f..lycoFen synthetase and

12.. 1°18 17.--ivc) •

Vasopressin has been observed to cause hepatic glycogenolysis

at a minimal effective concentration. of the same order as that necessary

for glucagon stimulation of glycogen breakdown. Since glucagon

stimulates glycogen phosphorylase under these conditions, the effects

of vasopressin and other glycogenolytic hormones on glycogen synthetase

and phosphorylase were evaluated.

Since glucagon is known to have a rapid, direct hepatic action

the response of the enzymes of glycogen metabolism to the hormones were the

assessed by injection into/hepatic portal vein of an intact animal (see

Methods, Section 2.4.2). Vasopressin caused a small but significant

increase in phosphorylase (Fig. 20) although the effect was very short-

Table 13. Effect of 8-ar -vaso•ressin. on he•atic cozen s thesis in vivo

Glucose (1.5M) was infused intravenously (3m1/h) into anaesthetised 48h-starved rats and

two liver samples were removed sequentially from each rat. At the time of the second liver sample, the

average blood glucose concentration was 33mM. Result are the mean of six experiments, ± S.E.M.

Additions to infusion fluid Glycogen content Oz mol of glucose / g) Calculated rate of glycogen synthesis ( u mol of glucose

/g/min) After 10min. After 70min.

None 35 72 0.62+0.09

Vasopressin 32

44

0.19+0.11 (175m units/ml)

124

I I I I 2

TIME NM)

Fig .20. - The time course of has hor lase activation by

Llycogenolytic hormones.

Experimental details are as in the text. Hormones (giucagon

1.0ug,n; adrenalin, 1.5 x 10 -8 mol, 0 ; (8-arginine)-vasopressin,

10 m units, A or 100 m units, A ) were injected into the hepatic

portal vein and the liver removed after various times, control,(0).

Phesphorylase was measured in crude liver homogenates. Results are

mean values + S.E.M. (indicated by bars) of at least 3 experiments,

except for 100 m units vasopressin at 3 and 5 min : one observation each,

and 10 m units vasopressin at 0.5min (mean of 2 values : 11.65 and 11.95).

125

tern, reaching a peak after 30sec. A similar elevation was observed with

adrenalin. The glucagon effect was slower than the other two hormones

but much more prolonged and the actual increase in phosphorylase was much

greater.

None of the hormones tested had a significant effect on glycogen

synthetase (Table 14), considered either as actual activity of the Tian

form of the enzyme or as the percentage of the enzyme in the "a" form.

126

Table 14. Effect of 1,co enol tic hormones on the activities of

glycogen synthetase

Experimental details are as in the texband Jig. 19.

Glycogen synthetase was measured essentially as Method 1, described in

the Methods Section (2.5.4), except for the assay being carried out in

the crude liver homogenate. Results are means ± S.E.Ms with the number

of observations in parentheses. Mean percentages of synthetase "a" were

calculated from ro "a" values in individual samples.

Injection Time after injection (min)

n a_ G1 co thetase

"a" Total "a"

None 0 6 0.15 + 0.02 0.49 + 0.04 31 ± 3

nazi 0.5 3 0.20 ± 0.04 0.58 + 0.04 35 + 4

1 3 0.18 + 0.01 0.63 + 0.02 28+ 1

3 3 0.16 + 0.02 0.40 + 0.02 40 + 5

5 4 0.13 + 0.03 0.35 4- 0.01 36 ± 8

(8-arginine) vasopressin

0.5 2 0.18 0.47 38

10 units 1 3 0.14 + 0.01 0.54 + 0.02 27 + 1

3 4 0.11 + 0.02 0.35 + 0.05 30 + 5

5 4 0.16 ± 0.01 0.40 + 0.04 40 ± 4

(8-arginine)- vasopressin

0.5 4 0.18 + 0.01 0.56 + 0.04 32 ± 3

100 units 1 3 0.16 + 0.05 0.42 + 0.04 37 ± 5

3 1 0.12 0.52 '23

5 1 0.16 0.60 26

Glucagon 1 3 0.16 4. 0.01 0.49 4- 0.06 34 + 3 1.0 lug

3 3 0.24 + 0.02 0.50 + 0.09 49 + 5

5 3 0.16 + 0.01 0.39 + 0.04 43 + 4

10 1 0.14 0.36 38

Adrenalin 0.5 4 0.13 4 0.01 0.41 ± 0.05 32 + 4 8mol 1.5x10 _

1 3 0.14 + 0.01 0.30 4- 0.04 47 + 3

2 4 0.15 + 0.02 0.38 + 0.06 43 + 4

3 3 0.14 + 0.01 0.31 4. 0.02 46 + 1

5 4 0.13 + 0.02 0.33 + 0.08 44 + 6

127

3.4 HEPATIC GLYCOGEN METABOLISM IN THE STARVED

STREPTOZOTOCIN-DIABETIC RAT

1. Glycogen accumulation in the. perfused liver

from diabetic rats.

Glycogen synthetase and phosphorylase

activities in vivo and in the perfused liver

of diabetic rats.

3. Influence of glucose and fructose on the

activities of glycogen synthetase and

phosphorylase in vivo.

Hepatic glycogen accumulation in the intact

diabetic rat.

128

3.4 HEPATIC GLYCOGEN METABOLISM IN THE STARVED STREPTOZOTOCIN

-DIABETIC RAT

As decribed above, net glycogen accumulation has been observed

in the perfused liver from starved rats. Under optimal conditions,

insulin added in vitro had no effect but in suboptimal conditions (i.e.,

glucose less than 30mM or in the absence of gluConeogenic precursors)

a moderate stimulation .of synthesis was observed (see Section 3.1.3),

The effect of insulin in hepatic synthesis warranted further study as,

for a variety of reasons the role of insulin in hepatic carbohydrate

metabolism is uncertain (see Introduction, Section 1.4.1). Therefore,

since conditions had. been established forst-udyingnet glycogen accumulation,

glycogen synthesis and the role of insulin in the process were studied

inn-the streptozotocin-diabetic animal.

3.4.1 Glycogen accumulation in the perfused liver from

diabetic rats

In the livers of starved-diabetic rats, there was a marked

decrease in rates of net glycogen accumulation compared to rates in

starved normal rats (Fig. 21; Compare Tables 11 and 15),'in which a linear

time course of glycogen deposition was maintained for at least 80min

(Fig. 21). This defect in glycogen synthesis in the livers of diabetic

rats, observed in perfusions with glucose plus either fructose or a

mixture of C3-substrates (lactate, glycerol and pyruvate), was completely

reversed by the administration in vivo of a mixture of glucose and fructose

50min prior to perfusion (Table 15). Glucose alone (in a dose equal to

that of the glucose & fructose together, i.e., 3mmols) was not effective

in correcting the impaired synthesis, and neither was fructose (0.5mmol)

alone (Table 15). Insulin treatment for similar periods (50 or 75min)

0 60

0

11.1 0 o 40

?-1 0

ce

20

129

Fig. 21

20 60

100

TIME (MIN)

Tiniecoirseofneizlx2aeaEj.a.ccumulation in normal and

diabetic rats.

Livers were perfused with bicarbonate-buffered saline containing

erythrocytes, and additions as indicated. Two samples (median followed

by left lateral lobe), or three samples (above two preceded by.right

lobe) were removed in sequence. Pilled-in symbols represent normal

(starved) rat livers, and open symbols diabetic rat livers. Additions

were as follows: (i) glucose (30mM) plus C3-substrates (.4 , 0 , 6

perfusions; M , 12 perfusions ); (ii) supplemented medium, including

insulin, hydrocortisone and amino acids, plus glucose, 30mM (0, five

perfusions; 0, two, perfusions), or glucose (20mM initially) and

fructose, 5mM ( v , 4 perfusions). Other details are given in the

text. Results are mean values, and bars indicate the S.E.M.

Table 15. Glycogen accumulation during perfusion of livers from streptozotocin-diabetic rats

Livers from 48h-starved diabetic rats were perfused for 50min with bicarbonate-albumin-saline containing washed erythrocytes. Substrates were added after 15min. Glucose was initially 28mM, and the C3-substrates, lactate, glycerol and pyruvate, were initially 5mM, 3.3mM and 1.3mM (respectively),thei.infused. Fructose was initially 5mM, and then infused (0.5M73m01). Pretreatments were administered subcutaneously, 50min before perfusion (unless indicated): glucose plus fructose in 2m1 , (1.25M and 1.25M respectively)7fructose (2m1) 0.25M) or glucose (2m17 1.5M). Other details are in the text. Results are mean +

Glycogen content Cumol of Calculated rate glucose per g of fresh liver) of glycogen

accumulation

Glucose concentration

(11`1)

Perfusions with Flucose,

20min (median lobe)

50min (left lateral

lobe) lactate, glycerol and pyruvate

Pretreatment No of perfusions

None 6 3.5 9.3

Glucose, fructose 6 48.1 72.3

Glucose, fructose & anti-insulin serum 4 6.9 18.6 Glucose, fructose & control serum 4 18.6 39.1

Insulin (50min) 5 11.3 26.6 Insulin (75min) 3 13.0 31.6 Perfusions with glucose plus fructose


None 7 7.1 17.7 Fructose 3 3.5 9.6 Glucose 3 42.9 54.3 Glucose, fructose 9 25.6 50.8

(jumol glucose/g/min) 20min 50min

0.19 + 0.04 27.1 + 0.9 28.7 + 1.3

0.81 + 0.12 29.4 + 1.2 32.0 + 1.4

0.39 4. 0.08 29.3 + 1.0 31.5 + 1.1

0.68 + 0.18 28.2 + 1.3 29.9 + 1.6

0.51 + 0.15 27.5 ± 0.8 29.0 + 0.9

0.62 4. 0.24 28.7 + 0.9 30.9 + 0.6

0.35 + 0.07 --- 25.6 ± 0.9 28.5 ± 1.1 0.20 + 0.06 30.4 + 1.6 33.7 + 1.3 0.38 + o.o5 30.6 ± 0.9 32.7 ± 0.9 H 0.84 + 0.13 31.0 4. 0.8 33.6 + 1.2 0

131

largely restored the rate of glycogen accumulation (Table 15).

The possibility that insulin was implicated in the response

to hexose pretreatment in vivo was investigated by administering anti-

insulin serum with the glucose and fructose; reduced rates of glycogen

synthesis were observed in the perfused liver compared to those after

treatment in vivo with glucose, fructose and control serum (Table 15).

The rate was not, however, reduced to that seen when there was no

treatment in vivo and the livers were perfused with C3-substrates. The

rate of glycogen accumulation, when fructose was present in the perfusion

medium, was higher than when lactate, glycerol and pyruvate were added,

as was also observed in normal (starved) livers (Section 3.1.5).

Insulin in the medium (in the absence of treatment in vivo) did not

increase the rate of glycogen synthesis from glucose plus fructose or

other substrates (Fig. 21., Table 17).

In an attempt to increase glycogen synthetic rates during

perfusion of livers from diabetic rats, the standard Medium was

supplemented with further substrates and hormones. Two groups of

perfusions were carried out. In one group, events during pretreatment

with glucose and fructose were simulated in the initial phase (about Ih)-

of perfusion : the medium contained fructose (initially 5E11), glucose

(20mM), and insulin, cortisol and amino acids (the latter three infused,

essentially according to John and Miller (1969)). After 50min , more

glucose was added to bring the concentration to about 30mM (optimal for

glycogen synthesis; see Section 3.1.2) and 03-substrates were added

(total 10111M initially, and then infused). Net glycogen accumulation

under these conditions, determined between 70 and 100min (Fig.21) was

no faster than in perfused livers of other -Untreated diabetic rats.

132

Thus fructose, glucose and insulin were not effective in restoring

glycogen synthesis when added to the perfusate.

In a second group of perfusions, the medium was supplemented

with insulin, hydrocortisone and amino acids (as above, but without

fructose) in addition to the standard condition of glucose (30mM) plus

C3-substrates. Liver samples were removed after 20 and 50min. There

was no improvement in the low rates of glycogen accumulation (Ilg. 21).

When the perfusion time was extended to 80min similar low rates of

glycogenesis were observed.

In control experiments, livers from normal (starved) rats

were perfused with thesupplemented medium; net glycogen synthesis (Fig.

21) occurred at a rate similar to that in perfusion; with glucose plus

C3-substrates (Fig.21 and Table 11) or fructose (Table 11). Thus,

supplementation did not inhibit synthesis in normal (starved) livers.

In all conditions, there was net output of glucose during

perfusion, as in livers of normal starved rats perfused in these

conditions (even during maximal glycogen deposition). If no glucose

was added to perfusions of diabetic rat livers, there was no glycogen

accumulation, and the rate of net gluconeogenesis (from C3-substrates)

was 1.11 0.20 (3))umol glucose/min/g of fresh liver (mean ± S.E.M.).

Thus the capacity for glucose synthesis was sufficient to support normal

net glycogen deposition (0.68 - 0.82)amol/min ,/g).

133

3.4.2 Glycogen synthetase and_philluhcmrlase activities

in vivo and in the perfused liver of diabetic

rats.

To gain insight into the impairment of net glycogen

accumulation in the perfused liver of diabetic rats the activities of

the enzymes glycogen synthetase and phosphorylase were measured.

Zacozmamthetase and phosphorylase activities in vivo.

The activities of glycogen synthetase and phosphorylase were

compared in intact normal and streptozotocin-diabetic rats (Table 16).

In starved diabetic rats, compared to normal starved rats, there was

a moderate decrease in the proportion of synthetase "a" due to an

increase in total activityl and an associated slight increase in

phosphorylase (expressed per g). These enzymes were assayed simultaneously

in the different lobes of the liver, as a pre-requisite to the

interpretation of their activities in lobes removed in sequence during

perfusion. There were no significant differences in the enzyme activites

between the various lobes, which allowed the enzymes and glycogen to be

measured in the same samples and the differences between sequential

samples to be assessed.

aycoreta.aeancl.ppahosholaseac'tyisia;the_pszf:used liver.

The enzymes glycogen synthetase and phosphorylase were assayed

in two sequentially removed samples during perfusion. After 50min perfusion

with glucose plus either fructose or C3-substrates (unsupplemented medium)

the proportion of glycogen synthetase in the "a" form (Method 1) was

51-60 (Table 17). In contrast to the result in normal (starved) rat

livers (Section 3.1.5) there was no clear correlation between glycogen

synthetase "a" activity (at 50min) and the rate of glycogen accumulation.

In most conditions the percentage of synthetase in the "a" form in the

initial sample (removed after 20min perfusion) was 23-40 which resembled

that in the intact diabetic animal (cf. Tables 16&17). In two groups of

Table 16. Activities of glycogen synthetase and phosphorylase in intact starved normal and starved

streptozotocin-diabetic rats.

Enzyme activities in simultaneously-sampled lobes of livers from 48h-starved rats were

assayed as described in the text. Results are means + S.E.M. from three normal and four diabetic rats. Mean /were

percentages of synthetase "a' calculated from % "a" values, in individual samples.

Glycogen synthetase (p.mol/imin/g : Method 1)

Glycogen phosphorylase (ipmol/min/g : centrifuged

Rat Lobe homogenate) "a" Total yo "a"

Normal Median 0.10 + 0.01 0.22 + 0.06 49 + 6 5.8 + 1.4

Normal Left lateral 0.13 + 0.01 0.32 + 0.05 45 4. 7 5.8 + 1.2

Normal Right 0.09 ± 0.02 0.20 + 0.04 43 3 4.5 + 0.7

Diabetic Median 0.14 + 0.01 0.37 + 0.01 39 + 5 6.1 + 0.8

Diabetic Left lateral 0.11 4- 0.02 0.33 + 0.02 34 + 8 6.4 + 0.6

Diabetic Right 0.13 + 0.04 0.35 + 0.07 38 ± 6 6.5 ± 0.3

Table 17. G1 cozen s thetase activi in the erfused liver of starved stre tozotocin-diabetic rats

Livers were perfused as described in Table 15., with various substrates and after different pretreatments.Glycogen synthetase was assayed (Method 1) in liver samples removed after 20 and 50 min. Details are in the text. Other results from these perfusions are included in Table 15. Results are means +

Glycogen synthetase (paolboin/g of fresh liver)

"a" Total % via!?

Calculated rate of glycogen accumulation

(pniol of glucose/ min per g)

20 min 50 min Perfusions with glucose, lactate, glycerol and pyruvate

"a" Total % "a"


None 4 0.13+0.01 0.30+0.05 48+6 0.11+0.02 0.19+0.03 60j3 0.17+0.09 Glucose, fructose 4 0.13+0.02 0.38+0.07 35+6 0.16+0.01 0.27+0.03 64-19 0.86+0.17 Glucose, fructose& anti-insulin serum

4. 0.14+0.15 0.32+0.05 461.5 0.14+0.02 0.25+0.06 61±9 0.39+0.08

Glucose, fructose & 4 control serum 0.10+0.03 0.23+0.03 42+8 0.09+0.01 0.16+0.01 56±9 0.68+0.18

Insulin 4 0.13+0.01 0.29+0.02 46+2 0.15+0.04 0.23+0.05 64+6 0.59+0.16 Nbne * 3 0.16+0.04 0.23+0.04 68+6 0.17+0.04 0.22+0.05 78+4 0.13+0.02 Perfusions with glucose, plus fructose


None 4 0.141:0.02 0.431-0.04 34±5 0.181-0.06 0.25-10.05 681-9 0.38+0.10 Fructose 2 0.09 0.33 28 0.09 0.18 51 0.19 Glucose 3 0.12±0.01. 0.33±0.06 39±7 0.13-10.02 0.20±0.01 66±15 0.38+0.05

.Glucose,fructose 4 0.16+0.04 0.2 650 0.121-0.01 0.19-10.02 64±5 0.86+0.21

* Supplemented medium, including insulin: samples at 70 and 100 min.: See Fig.20

136

perfusions, where the liver was in contact with fructose for about

100min (in vivo and or perfusion), the percentage of synthetase "a"

was higher (Table 17) : (i) pretreatment and perfusion with glucose and

fructose (65 and 64% "a"), (ii) no pretreatment in vivo, but prolonged

perfusion with glucose and fructose in supplemented medium (68.and 78%

"a"). These high values for synthetase "a" were associated with rapid

glycogen accumulation in the former situation, but not the latter.

The activities•of glycogen phosphorylase in liver samples

removed during perfusion were lower than those in intact diabetic rats

(cf. Tables 16 & 18). The activities at 20 or 50min did not conform

to any pattern (Table 18). Changes in glycogen synthetase and

phosphorylase during perfusion were assessed from the activities in

sequential liver samples. In the livers of diabetic rats,when net

glycogen accumulation was restored by pretreatment,there was an increase

in the proportion of glycogen synthetase "a" during perfusion (Table 17),

which presumably reflected the activating effect of substrates on

synthetase, as observed in perfused livers of normal (starved) rats

(Section 3.1.5). This response was much less marked in diabetic perfusions

where glycogen synthesis was low (Table 17). Taking all groups of

diabetic perfusions with glucose and C3-substrates, the increase in the

proportion of synthetase "a" between 20 and 50min was correlated with

the rate of net glycogen accumulation (Pig 22a). In perfusions with

fructose there was no such correlation (Table 17).

In the experiments described above, the alteration in %

synthetase "a" during perfusion was manifested by a fall in total enzyme

(Table 17) rather than in the absolute value of synthetase "a". This

may have reflected alteration in the distribution of synthetase in

Table 18. Phosphorylase activity in the perfused liver of starved streptozotocin-diabetic rats

Livers were perfused as described in Table 17 (and Table 15). Glycogen phosphorylase was

assayed in (centrifuged) samples for which the glycogen synthetase results are given in Table 17. Other details

are in Table 17, or the text.

Glycogen phosphorylase Rate of glycogen (Fmol/Min/g of fresh liver) accumulation

!amo1 glucose / 20min 50min min per g)

Perfusions with lucose, lactate,, glycerol and pyruvate

Pretreatment:

None 1.93 + 0.48 1.40 + 0.23 0.17

Glucose, fructose 2.92 ± 0.58 1.72 + 0.52 0.86 \

Glucose, fructose,anti -insulin serum 3.12 ± 0.51 2.53 ± 0.75 0.39

Glucose, fructose, control serum 1.78 ± 0.23 1.46 ± 0.24 0.68

Insulin 2.23 + 0.52 1.25 ± 0.24 0.59

None * 2.62 ± 0.87 2.34 ± 0.52 0.13

'Perfusions with glucose plus fructose

Pretreatment:

None 2.55 + 0.06 1.38 + 0.21 0.38

Fructose 2.09 1.47 0.19

Glucose 1.72 + 0.53 1.51 + 0.47 0.38 H

Glucose plus fructose 2,39 + 0.25 1.48 + 0.20 0.86 ■' 4

* Supplemented medium, samples at 70 and 100 min: See Fig.21

•

_ 138

For Figure 22. See Over—leaf

Fig. 22. Relationship between net hepatic glycogen

accumulation and glycogen synthetase and

phosphorylase in diabetic rats.

In livers perfused with glucose plus 03-

substrates, for which glycogen changes are given in Table 15,

and measurements of glycogen synthetase and phosphorylase

activities (pmol/g/min) in Tables 17 and 18, mean values for

the change between 20 and 50min perfusion in (a) the

percentage of glycogen synthetase "a" and (b) the quotient

(percentage synthetase "a")/(phosphorylase) were calculated.

Rats received either no pre-treatment (o, no additions; A 7

supplemented medium; see Fig. 21) or insulin ( A ), or glucose

and fructose (o ), or hexoses plus either anti-insulin serum

( a ) or control serum ( 0 )0 Other details are in the text.

Bars indicate S.E.M., and the numbers of perfusions are in

Table 17. Lines were fitted by regression analysis of values

from individual perfusions : (a) r = 0.54, p < 0.01

(23 observations); (b) r = 0.64, p < 0.01 (23 observations).

0

O

3 0 CD

C0 co

3 0

CD

•;•• c0

<A

tn

O

O co

O

CHANGE IN ENZYME ACTIVITY IN 30 MIN PERFUSION

Increase in (% synthetase"al(phosphorylase) Increase in % glycogen synthetase "a" O 0 11 8 0

140

centrifuged homogenates, or could have been a consequence of the

action of modifiers in the homogenates if UDPG was not at saturating

concentrations (see discussions by Bishop and Larner, 1967; Blatt and

Kim, 1971b). To distinguish between these possibilities the details

of the synthetase assay were altered (Method 2) and the enzyme measured

in key groups of perfusions. In these uncentrifuged homogenates, there

was no decline in total synthetase during perfusion (Table 19). However,

the main finding of the experiments using Method 1 were confirmed, i.e.,

the proportion of synthetase "a" (and the absolute value) increased

duririg perfusions of livers from diabetic pretreated rats, but not in

livers of un -treated animals (Table 19). The rates of glycogen deposition

resembled those in Table 15; with insulin in the perfusate, the average

rate was 0.26pmol of glucose/Min/g measured in three perfusions, confirming

the lack of a direct hepatic action.

There was a decline in the activity of phosphorylase (assessed

between 20 and 50min) during perfusion of diabetic rat livers (Table 18)

as would be expected as glycogen synthetase and phosphorylase often

exhibit reciprocal changes. The extent of the changes in each enzyme

were correlated (Fig. 23). There was relatively little decline in

phosphorylase during perfusion of liver from diabetic rats with glucose

plus 03-substrates (0.53 + 0.26 (4) pmol/min/g fresh liver) compared

to thegreaterdecrease in perfusions after pre-treatment in vivo (1.20

0.38 (4) jumol/min/g), in which net glycogen accumulation was more rapid

(Table 18). Taking all groups, there was not however, any clear-cut

inverse correlation between the net rate of glycogen synthesis and the

decline in phosphorylase activity during perfusions (with or without

fructose in the perfusate).

Table 19. Glycogen synthetase activity in the perfused liver of starved streptozotocin-

diabetic rats.

Livers were perfused as described in Tables 15 and 17, with glucose, lactate, glycerol

and pyruvate, and glycogen synthetase was assayed (Method 2). In one group (*) insulin was infused into the

perfusion medium: 1.5m1A of a solution containing 330mU/M1., following an intial dose of 500mU. Other details

are in the text. Results are means +

Glycogen synthetase cumol/min/g of fresh liver : Method 2)

Pretreatment (in vivo). No of perfusions.

20 min 50 min

"a" Total % "a" "tar" Total % nal?

None 0.52 + 0.09 0.84 + 0.11 62 + 7 0.55 + 0.40 1.05 + 0.12 53 4. 4

Glucose, fructose 3 0.64 + 0.05 0.98 + 0.09 66 ± 6 0.83 + 0.10 1.11 + 0.08 .75 + 6

Insulin (IU, 75 min, before perfusion) 4 0.60 + 0.16 1.12 + 0.23 52 + 3 0.78 + 0.15 1.12 + 0.18 69 + 5

None * 6 0.51 + 0.07 0.99 + 0.07 51 + 6 0.52 + 0.09 0.99 ± 0.08 52 + 6

* Insulin added to perfusion medium

0 A

z 0 L.

w

z

C)

z-

CH

AN

GE

1 N

GLY

CO

GE N SYN

TH

ETASE "a

"

+20

0

142

•1.40

0

+0.4 0 -OA -0.8 -1.2 -2.0

CHANGE IN PHOSPHORYLASE ACTIVITY IN 30 MIN PERFUSION

Fig. 23. Relationship between the changes in glycogen synthetase

and phosphorylase during liver perfusion in diabetic rats

Livers were perfused with glucose plus 03-substrates, with or

without pre-treatment in vivo (see Tables 15, 17 and 18, Pig. 22). an Chges in the activities of glycogen synthetase (percentage "a" and

phosphorylase Oa mol/min/g) between 20 and 50min are presented for the individual perfusions described in Tables 17 and 18. Symbols are as in Fig. 22. Other details are in the text. The line was fitted by regression analysis, taking into account variation in both axes : r = 0.67, p < 0.001. (23 observations).

143

The above results demonstrated that the response of enzymes

to glucose and 03-substrates during perfusion was impaired in diabetic

rat livers, anoVres was tored after pretreatment in vivo with hexoses or

insulin. A combined measure of the functional state of both synthetase

and phosphorylase is the quotient (% synthetase "a")/(phosphorylase). The

change in this quotient between 20 and 50min perfusion with glucose plus

C3-substrates was significantly correlated with the net rate of glycogen

accumulation in all groups if diabetic rat livers (Fig. 22b).

3.4.3. Influence of glucose

of glycogen synthetase andhosse in vivo

The impairment in glycogen accumulation observed in the

perfused liver of diabetic rats was reversed by the administration in

vivo of a mixture of glucose and fructose 50min prior to perfusion (see

Section 3.4.1),It was therefore of interest to study the effect of this

treatment in vivo on glycogen synthetase and phosphorylase.

In normal rats, glucose and fructose each induced conversion

of synthetase to the "a" form (Table 20). Glucose and fructose together

brought about the greatest "activation" of the enzyme (Table 20) in

agreement with observations in the perfused liver (Table 11; Section

3.1.5). In diabetic rats these effects were less marked (Table 20).

Administration of the hexoses, either alone or in combination, also

produced a decline in phosphorylase activity in normal rats (Table 20),

although the effect of glucose plus fructose was not greater than either

hexose alone. A similar response was seen in the diabetic rat indicating

that control of phosphorylase is not impaired.

Table 20. Influence of :lucose and fructose on lons thetase and •hosh lase in

intact starved normal and streptozotocin-diabetic rats.

Enzyme activities in simultaneously-sampled lobes from 48h-starved rat livers were

assayed as described in the text. - Treatments were administered in 2m1, S.C., 50 min before sampling: glucose

(2.5M), fructose (0.25M) or glucose plus fructose (1.25M and 0.25M respectively). For comparison, two values

from Table 16 are included. Other details

all groups except fructoss-treated diabetic

are in the text. Results are means

rats (5 animals).

Glycogen synthetase (% "a"). (Method 1)

± S.E.M. from three rats, in

Glycogen phosphorylase centrifuged homogenate) pmol min g)

Rat Treatment Median lobe Left lateral lobe Median lobe Left lateral

lobe Normal -* 49 + 6 45 + 7 5.8 4- 1.4 5.1 4. 1.2

Normal Glucose 77 + 2 70 + 4 2.1 + 0.3 1.5 + 0.2

Normal Fructose 60 ± 5 69 .... 4 2.7 + 0.4 2.1 ± 0.2

Normal Glucose plus fructose 83 + 7 81.- 6 2.3 + 0.1 2.7 + 0.2

Diabetic -* 39 + 5 34 + 8 6.1 + 0.8 6.4 ± 0.6

Diabetic Glucose 50 ± 5 57 + 7 2.6 + 0.4 2.6 4. 0.1

Diabetic Fructose 46 ± 7 46 + 6 3.5 ± 0.6 2.5 ± 0.4

Diabetic Glucose plus fructose 47 4- 6 48 ± 3 3.7.4- 0.3 3.1 4. 0.6

* From Table 16 for comparison 1-,

145

3.4.4 Hepatic glyco gen accumulation in the intact

diabetic (starved). rat.

For comparison with resultsin the perfused liver, glycogen

accumulation was followed in anaesthetised intact streptozotocin-

diabetic rats. An impairment in the capacity for net glycogen synthesis

was observed during infusion of either glucose or glucose plus a mixture

of glycerol, serine and pyruvate, following pre treatment in vivo

(Table 21). As in the experiments with the perfused liver, fructose

and glucose were required to bring about rapid glycogen deposition

in diabetic rats (Table 21).

Table 21. Glycogen accumulation in intact starved normal and starved streptozotocin-diabetic rats.

Glycogen accumulation was followed in intact rats. Rats received various pre-treatments (S.C.,1°

in two portions of Iml , unless indicated; see Table 15), were then anaesthetised with Nembutal, and received

an infusion (3ml/h) into a tail vein of glucose (1-1.25K), fructose (0.25K) or gluconeogenic C3-substrates (a

mixture of glycerol, serine and pyruvate, each 0.33M, also used for pre-treatment). The first liver sample was

removed within about 15min of anaesthesia, and the second lh later. The indicated time of pre-treatment refers

to the interval between injection and the first sample. Other details are in the text. Results are means S.E.M.

Rats Pre-Treatment Infusion Glycogen content ()umol

No. of of glucose per g of experiments fresh liver

Sample 1 Sample 2


ol glucose,/ g/mmn)

Final blood glucose concentration

(lox)

Glucose

Glucose plus fructose

Glucose

Glucose

Glucose plus C3 -substrates



Normal None

Normal Glucose plus fructose (60min)

Diabetic Glucose 12min)

Diabetic Glucose (53min)

Diabetic Glucose plus C -substrates 7

Diabetic Glucose plus fructose (i.g., 35min)

Diabetic Glucose plus fructose (60min)

35.2 72.3 0.62 4. 0.09 32±4 4 110.6 155.0 0.74 4. 0.15 36 4. 7

2 5.2* 8.4* 0.06 25

3 77.2 70.6 0 37 ± 5 3 11.5* 17.9* 0.12 4. 0.10 64 ± 17

4 26.6 63.2 0.61 4. 0.18 61+9

72.1 109.9 0.63 ± 0.08 40 ± 3

(i.g.,

* Samples 53min. apart Abbreviations : s.c., subcutaneously; i.g., intragastrically.

rn

147

3.5 HEPATIC CARBOHYDRATE METABOLISM IN THE STARVED

ADRENALECTOMISED RAT

1. Glycogen accumulation in the perfused

liver from adrenalectomised rats.

2. Glycogen synthetase and phosphorylase

activities in vivo and in the

perfused liver of adrenalectomised

rats.

148

3.5 HEPATIC CARBOHYDRATE METABOLISM IN THE STARVED

ADRENATRCTOMISED RAT

Net rates of glycogen accumulation were obtained in the

perfused liver of normal starved (see Section 3.1) and diabetic

starved rats (see Section 3.4), and the relationships established

between glycogen and the enzymes of its metabolism, glycogen synthetase

and phosphorylase. Since the role of adrenal cortical steroids in

glycogen metabolism is not clear (see Introduction, Section 1.4.2)

it was of interest to study glycogenesis and its control in the et

adrenalectomised rat under conditions optimal fo rates of glycogen

accumulation in other states.

3.5.1 Glycogen accumulation in the_perfused

liver from adrenalectomised rats

Glycogen synthesis was studied in the perfused liver from

starved adrenalectomised rats in the conditions which allowed net

rates of accumulation in the normal starved rat; no synthesis was

observed (Table 22). Supplementation of the perfusion medium with

hydrocortisone, amino acids and insulin did not increase the rate of

glycogensis. Pre-treatment of the rat in vivo for 50min with glucose

and fructose prior to perfusion of the liver, (a treatment which restored

synthesis in the diabetic rats : see Section 3.4.1) increased synthesis

in the presence or absence of hydrocortisone or insulin in vitro,

but not to the rates observed (Table 22) in sham-operated rats. In

all these perfusions glucose concentration was 27.5-30.5mM. There was

slight glucose output during perfusion of sham-operated or pre-treated

adrenalectomised rats (Table 22). Thus the net carbon source of

glycogen in these perfusions was C3-substrates, rather than glucose,

as in similar perfusions of starved (Section 3.1), or starved diabetic

(Section 3.4) rat livers. In livers from untreated adrenalectomised

rats (where there was no glycogen synthesis : Table 22), there was no

149

For Table 22, see over-leaf

Table 22 Glycogen accumulation in the perfused liver from adrenalectomised starved rail

Livers from 48h- starved adrenalectomised rats were perfused as described in the text.

In all perfusions, glucose was initially 28mM and3-substrates (lactate, glycerol and pyruvate) were

5mM, 3.3mM and 1.3m1v1 (respectively) and then infused at 3m1/h from 15min. The full supplement medium

containing insulin, amino acids and hydrocortisone was as described in the Methods (Section 2.3.2).

Pre--treatment was administered subcutaneously at the time indicated before perfusion : glucose plus

fructose in 2m1 (1025M and 0.25M respectively) and 1U of insulin. Results are means -I- S.E.M. with

the number of observations in parentheses.

Livers were perfused with glucose (28mM), fructose (5mM) and insulin (added

and infused) for 2h,3-substrates were added and infused with insulin from 135min and liver and medium

samples were taken at 140 and 170min.

Livers were perfused with glucose (28mM), fructose (5mM), C3-substrates (10mM),

amino acids mixture (4 x "normal"), hydrocortisone and insulin (added and infused). After 60min perfusion,

amino acids (2 x "normal"), fructose (5mM) and 03-substrates (10mM) were added. The first and latter of

these were also added after 120min perfusion. C3-substrates were infused from 165min, liver biopsies and

medium samples being removed at 180 and 240min.

Sham-operated controls.

208+1.5 2,1+1.4 27.5+0.8 26.6+0.9 -0.02+0.02 (6)

Full supplement

1.5+1.0 .1.7+0.6 28.5+0.6 28.9+0.7 0.01+0.04 (5)

2.8+1.2 7.4+1.8 29.1+1.2 32.0+1.5 0.16+0.05 (6)

Full supplement

5.8+2.0 15.1+4.3 27.5+1.1 28.1+1.0 0.31+0.09 (9)

Insulin 6.0+1.4 10.5:0.9 29.4+1.9 30.9+1.6 0.15+0.13 (3)

Hydro-cortisone

3.2+1.0 9.6+1.5 28.3+1.3 30.8+1.2 0.21+0.05 (4)

Hydro-cortisone

6.0+4.8 14.4+6.7 28.3+1.2 30.0+0.9 0.28+0.09 (s)

Hydro- cortisone &

1.6+0.5 9.0±3.4 26.6+0.9 28.4+1.1 .0025+0.10 (6)

Insulin

*Insulin & fructose

*0.6+0.1 *1.5+0.5 *32.7+0.7 *32.9+0.9 0.03+0.01 (3)

+Fructose & +18.2 +14.2 +32.5 1-33.3 -0.07 (2) Full supplement

19.3+4.9 38.8+6.0 28.4+0.7 50.4+0.9 0.65+0.06 (11)

Fructose & glucose

Fructose &

50 glucose

Fructose & gltcose

Fructose &

50 glucose

Fructose, 60 glucose & insulin



Glycogen content Pre-Treatment Time Additions to (rol of glucose/g fresh liver)

(min) perfusion 20min 50min 20min

(median lobe) (left lateral lobe)


50min (pmol glucose/g/min)

Glucose in medium (DE)

151

change in glucose concentration during perfusion. If glucose was

not added to such perfusions (with the mixture of lactate, glycerol

and pyruvate), the rate of net gluconeogenesis was about 1.5pmol/dmin (C.J

Kirk : unpublished results). Thus the capacity for gluconeogenesis

in these perfusions was sufficient to have supported glycogen synthesis.

In order to evaluate the impairment in glycogen accumulation

in the perfusion, adrenalectomisedrats were treated with hydrocortisone

in vivo for varying times prior to liver perfusion. Net rates of

glycogenesis (measured between 20 and 50min perfusion) comparable to

those observed in normal animals, were obtained when the animal was

pre-treated for 4h before perfusion (Fig. 24). When hexoses were

given with the steroid the restoration in glycogen accumulation was

seen earlier (Fig. 24) : significant rates being observed between 20

and 50 min of perfusion, after 2h pre-treatment in vivo.

The above time course of restoration of synthesis by

hydrocortisone suggested that net rates of glycogen accumulation were

not restored by the initial attempt at restoration with substrates,due

to the short pre-treatment time. Since it is likely that the hexose

treatment in vivo causes insulin secretion, which could have a role

in restoration, insulin was given in addition to the glucose and

fructose, and the rates of glycogenesis measured in perfusion after

varying times of pre-treatment in vivo. After 2h, in vivo treatment,

normal net rates of glycogen accumulation were obtained in the

perfusion (Fig. 24) which were not increased by longer treatment before

perfusion. Insulin in vitro had no effect on the rate of synthesis

after lh pre-treatment (cf. Table 22 & Fig. 24) although the addition

of hydrocortisone to the perfusion medium perhaps had a small effect

in the short -term (cf. Table 22 & Fig. 24).Due to this apparent

stimulation of hepatic glycogen accumulation by hydrocortisone in vitro

(during perfusion for 50min) the effect of this hormone in perfusion

was assessed over a longer time period. In the presence of amino acids,

LL. 0

w F-

GLY

CO

GEN

SYN

THES

IS

.152

0 1 • 2 3. 4 LENGTH OF PRETREATMENT PRIOR TO PERFUSION ( H

Fig. 24 Time course of restoration of net rates oflaa22gen.

position in the perfused liver from adrenalectbmised

rats.'

Rats were treated subcutaneously with fructose, glucose and insulin, ( as described in Table 22),o ; hydrocortisone succinate,

A 10mg; or fructose, glucose and hydrocortisone succinate (l.2517, 0.25M and 10mg, respectively)pyith the addition of insulin for lh value;or no pre-treatment, 0. After various times of pre-treatment of the rat, the liver was perfused with glucose (28mM) and C7-substrates as indicated in Table 22, and liver biopsies taken at 20 .ind 50min. Results are means of at least 3 experiments and bars indicate the S.E.M.

153

fructose, insulin and hydrocortisone the rate of glycogen synthesis

measured in liver biopsies removed after 3 and 4h perfusion was howevernegligible (Table 22).

Net rates of glycogen accumulation were therefore observed

in the perfused liver only after treatment in vivo with fructose,

glucose and insulin for 2h. When these in vivo events were simulated

in the perfusion and liver samples removed at 140 and 170min no

synthesis was obtained (Table 22).

For comparison, it may be mentioned that studies in the

intact anaethetised adrenalectomised rat have demonstrated that

hepatic glycogen accumulation is also impaired in vivo in short-term

experiments (C. Kirk, unpublished results). Thus when glucose was

infused for lh and liver biopsies removed for glycogen determination,

the rate of synthesis was negligible (about 0.03 pmol glucose/g/min)

in the adrenalectomised rat (cf. Table 21, Section 3.4.4). If

glucose was administered (sub-cutaneously or intra-gastrically) 1.5 -

2h before anaesthesia ratesof about one-third normal (0.1-0.37 iumoVe min) were observed on subsequent glucose infusion. These results

support (by a similar procedure involving sequential biopsies in

vivo) the find.irg that glycogen accumulation is impaired in the perfused

liver from adrenalectomised rats and that synthesis is not restored

in the short-term, by treatment with hexoses.

3.5.2 Glycogen synthetase and activities

in vivo and in the erfused liver of


As .a pre-requisite to the interpretation of the activities

of glycogen synthetase and phosphorylase in lobes of the liver

removed in sequence during perfusion, and to gain insight into the

impairment of net glycogen accumulation in the perfused liver, tie

enzyme activities were measured in the intact rat.

154

SycoaensynIhetase and hos hor lase activities in vivo

Glycogen synthetase.and phosphorylase activities were not significantly

different in the sham-operated animal and normal rat (Table 23, cf.

Table 16 Section 3.4.2). In the starved adrenalectomised rat the

activity of (supernatant) hepatic glycogen synthetase (total) was increased,

producing a significant decrease in the % "a" form (Table 23). The

relatively greater recovery of (total) synthetase in the high-speed

supernatant fraction in adrenalectomised rats (Table 23) may reflect

their lower glycogen content, i.e., there would be less "particulate"

synthetase. No difference between the various lobes of the liver in

either group of rats was observed; this allowed differences between

activities in sequential liver biopsies to be assessed as rates of change.

amogen svnthetase and phosBhozylase activities in the erfused liver.

The enzymes of glycogen metabolism were measured in two sequentially

removed samples during perfusion. As was noted in normal (starved) rat

livers (Table 11; Section 3.1.5) but not in diabetics (Table 17; Section

3.4.2) there was a correlation between the % "a" form at 50min and the

rate of glycogen accumulation (Table 24). This correlation was also

observed when fructose was substituted for C3-substrates in the perfusate,

unlike the results obtained in the diabetic perfusions (see Table 17,

Section 3.4.2). Since these assays were carried out in the uncentrifuged

homogenate, no loss of total enzyme activity during perfusion was recorded

(see also Section 3.4.2); thus a correlation between the actual activity

of the "a" form at 50min and rate of synthesis was also observed (Table

24). In most conditions the percentage of synthetase "a" in the initial

sample (removed after 20min perfusion) was 21-38% which was slightly greater

than that in the intact adrenalectomised rat (cf. Tables 24 & 23),

assuming that centrifugation did not decrease % "a" (see Section 3.1.4).

The activities of glycogen phosphorylase in liver samples

removed during perfusion were lower than those in the intact

Table 23. Glycogen synthetase and thosphorylase activities intact sham adrenalectomised

and adrenalectomised rats.

Enzyme activities in simultaneously-sampled lobes of livers from 48h-starved rats were

assayed as described in the text. Results are means + S.E.M. from three rats. % "a" was calculated for each

sample in order to obtain the S.E.M. for the mean values.

Glycogen synthetase (Method 1) Phosphorylase Rat Liver lobe (pnol / min g) (Centrifuged homogenate)

1•11.1■1111.1•1■011■1•••■■••••■•••■••••••••••••••• ••

"a" Total % n d "a" (pmol/minig)

Sham adx. Median 0.13 + 0.03 0.23 + 0.01 57 + 8 4.68 + 1.30

Sham adx. Left lateral 0.13 + 0.02 0.26 + 0.06 55 + 8 4.59 + 0.60

Sham adx. Right 0.13 + 0.03 0.28 + 0.01 47 + 7 5.40 + 0.24

Adx. Median 0.10 + 0.02 0.45 + 0.02 23 + 6 5.07 + 0.28

Adx. Left lateral 0.11 + 0.03 0.46 + 0.02 24 + 8 5.09 + 0.45

Adx. Right 0.09 + 0.02 0.47 + 0.04 21 + 6 5.01 + 0.53

Abbreviation : Adx. adrenalectomised.

Table 24. Glycogen synthetase activity in the perfused liver from starved adrenalectomised rats.

Livers were perfused as described in Table 22 and pre-treatment with F & G (fructose and glucose) was for 50min prior to perfusion. Glycogen synthetase was assayed (Method 1; uncentrifuged homogenate) in the same liver samples as was glycogen. Results are means + % "a" was calculated for each perfusion in order to obtain an estimate S.E.M. for the mean value.

Glycogen synthetase (pmol/Min/g) Rate of Pre-Treatment . Additions. to Rat No. of 20min L_50min glycogeh

the perfusion perfusions "a" Total % "a" ITan Total % "a" synthesis (Fmol glucose/ min/g)

Sham 3 0.34+0.11 0.56+0.12 56+11 0.53+0.09+ 0.67+0.05 79+8 0.61+0.08

- - Sham 3* 0.20+0.02 0.32+0.02 61./3 0.19+0.02 0.26+0.04 74+5 0.68+0.15

- - Adic. 3 0.12+0.01 0.48+0.03 250 0.14+0.01 0.55+0.05 25+2 -0.04+0.03

- *.13 Fructose Adx. 3 0.10+0.03 0.47+0.08 21+4 0.14+0.02 0.49+0.04 28+4 0.00+0.02

F & G - Adx. 3 0.14+0.05 0.52+0.10 26+5 0.26+0.05 0.66+0.06 38+4 0.19+0.10

F & G Full supple Adx. 4 0.21+0.04 0.53+0.03 38+6 0.35+0.08 0.73+0.02 47+11 0.31+0.17 -ment

- Full supple Adx. 3 0.13+0.01 0.43+0.05 31±3 0.12+0.01 0.44+0.05 270 -0.01+0.04 -ment

F & G Insulin Adx. 3 0.18+0.05 0.62+0.01 29+8 0.24+0.07 0.67+0.08 34+7 0.15+0.13

F & G Hydro- cortisone

Adx. 3 0.19+0.05 0.49+0.04 37+7 0.25+0.05 0.55+0.03 46±9 0.34+0.07

* Enzyme assayed in centrifuged homogenate (data not included in Fig. 25, 27 & 28)

..1' Fructose substituted for C3-substrates in perfusate

Abbreviations: adx. adrenalectomised sham. sham operated animal rn

F. fructose

G. glucose

157

adrenalectomised rat (cf. Tables 25 & 23), despite the fact that the

perfused liver samples were not centrifuged before assay. The activities

at 20 or 50min did not conform to any pattern (Table 25) as was observed

in the diabetic perfusions (Table 18; Section 3.4.2).

Changes in glycogen synthetase and phosphorylase during

perfusion were assessed from the activities in sequential liver samples.

When net rates of glycogen accumulation were observed in the livers of

sham-operated animals there was an increase in the percentage of glycogen

synthetase "a" during perfusion (Table 24). This response was less

marked in the adrenalectomised perfusions where glycogen synthesis was

low (Table 24). Taking all groups of perfusions, the increase in the

proportion of synthetase "a" between 20 and 50min (presumably reflecting

the enzyme activation by substrates) was correlated with the rate of net

glycogen accumulation (Fig. 25a). A similar enzyme sensitivity was

observed in the diabetic rat liver perfusions (see Fig. 22a; Section

3.4.2).

As has been noted previously (see Section 3.4.2) when the

glycogen synthetase activity was assayed in the centrifuged homogenate

the alteration in % "a" during perfusion was manifested by a fall in total

enzyme (Table 24) rather than in the absolute value of synthetase "a".

This fall was not observed when the crude liver homogenate was assayed,

when an increase in the absolute activity of the "a" form as well as the

percentage was seen during perfusion (Table 24). This increase in the

activity of the "a" form was correlated with the rate of glycogen

accumulation (Fig. 25b).

• When net rates of glycogen accumulation were measured in

the perfusion there was a decline in the activity of phosphorylase

(assessed between 20 and 50min); an increase occurred when there was a

negative or small rate of glycogen synthesis (Table 25). The change in

activity of phosphorylase between 20 and 50min was correlated with the

Table 25. Glycogen phosphorylase activity in the perfused liver from starved adrenalectomised rats.

All details are in Tables 24 & 22. Phosphorylase was assayed in Uncentrifuged homogenates

of the same liver samples as was glycogen synthetase and glycogen.

Phosphorylase ol/min/g)

Pre-Treatment Additions to the perfusion

Rat No. of perfusions 20min 50min

Sham 3 3.05+0.09 2.69+0.11

Sham 3* 2.31+0.40 1.65+0.11

Adx. 3 1.74+0.14 2.25-3.17

Fructose Adx. 3 2.26+0.52 3.16+0.73

F & G Adx. 3 1.76+0.0s 1.93+0.29

F & G Supple- mented

Adx. 4 2.32+0.07 2.49+0.10

Supple- mented

• Adx. 3 1.90+0.23 2.72+0.52

F & G Insulin Adx. 3 1.95+0.33 2.48+0.52

F & G Hydro- cortisone

Adx. 3 2.36+0.25 2.18+0.09

Rate of glycogen synthesis

Oumol glucose/Min/ g)

0.61+0.08

0.68+0.15

-0.04+0.03

0.00+0.02

0.19+0.10

0.31+0.17

-0.01+0.04

0.15+0.13

0.34+0.07

Fructose substituted for 03-substrates in perfusate

Enzyme assayed in centrifuged homogenate (data not included in Fig. 26)

Abbreviations : adx: adrenalectomised sham: sham operated animal Ft fructose G: glucose

For fig. 25 see Over-Leaf

159

Fig. 25 Change in sensitivity of glycogen synthetase

during net

in perfused livers from adrenalectomised rats.

In livers perfused with glucose plus C3-substrates,

for which glycogen changes are given in Table 22, and measure-

ments of synthetase in Table 24, mean values for the change

between 20 and 50min perfusion in (a) the percentage of glycogen

synthetase "a" and (b) the activity of glycogen synthetase "a"

(Ilmol/min/g) were calculated. Rats received either no pre-

treatment (0, no additions; A, supplemented medium : see Methods;

D , Sham-operated animals), or fructose and glucose (0, no

additions; A , supplemented medium; G3 , insulin; V , hydrocortisone).

Other details are in the text. Bars indicate S.E.M. and the

number of perfusions within each group are in Table 24. Lines

were fitted by regression analysis of values from individual

perfusions: (a) r = 0.72, p< 0.001 (22 observations);

(b) r = 0.73, p< 0.001 (22 observations).

Increase in % glycogen synthetase. "a"

0

❑ I

uo!p

alnw

mpo

a CD

0

2 - o

ca

3 o ca

CHANGE IN GLYCOGEN SYNTHET,ASE ACTIVITY N 30 .MIN PERFUSION

0

Increase in glycogen synth eta se "a" 0 0 O 0

(IA 1>--1

161

rate of glycogenesis (Fig. 26). A similar fall in phosphorylase

activity due to substrates in perfusate was observed during perfusion

of diabetic livers (see Section 3.4.2). Although the enzyme was assayed

- in the uncentrifuged homogenate of adrenalectomised rat livers the

decrease was less, for a corresponding rate of net glycogen accumulation,

i.e., a smaller decrease in phosphorylase activity was observed than in

the(centrifuged) samples from diabetic rat liver perfusions. As might

be expected, the alterations, during perfusion, in both glycogen

synthetase and phosphorylase were correlated (Fig. 27a & b).

The above results demonstrated that the response of enzymes

to glucose and 03-substrates during perfusion was impaired in adrenalect-

omised rat livers. A combined measure of the functional state of both

synthetase and phosphorylase is the quotient % glycogen synthetase "a"

/phosphorylase or glycogen synthetase "a" / phosphorylase. The change

in these quotients between 20 and 50min perfusion were significantly

correlated with the net rate of glycogen accumulation in all groups of

adrenalectomised rat livers (Fig. 28). When the results are compared

with those from diabetic rat liver perfusions (Fig. 22b, Section 3.4.2)

it would appear that the quotient % glycogen synthetase "a" / phosphorylase

did not increase by as much during adrenalectomised rat liver perfusions,

as was observed in the diabetic rat. As has been seen, the change in %

"a" during perfusion of adrenalectomised rat liver was similar to that

obtained in perfused diabetic rat livers. However, phosphorylase did

not appear to respond to substrates to the normal degree at agiven

glycogen synthesis rate : this was illustrated previously (Fig. 26) and

in the proportional changes of both enzymes during perfusion (cf. Fig 27 &

23),It would therefore seem that'in the adrenalectomised rat liver, the

response of glycogen phosphorylase to activitors or inhibitors is impaired,

as reflected in the lower change in activity, at a given glycogen synthetic

rate, compared to that in normal (or diabetic) rats.

0

IN 3

0 M

IN PE

RFU

SIO

N

+1. 0 162

.111■,•1

0. 4

+0.2

0.1 0.2 0.3 0 = 0.'5 0.6 0.7

•••••■11.

-0.6-- RATE OF NET GLYCOGEN ACCUMULATION (11MOLI GI MIN )

Fig. 26 Response of -phosphorylase activity during net rates sgely_ coon. accumulation.

In livers perfused with glucose plus 03-substrates, for which

glycogen changes are given in Table 22, and measurements of phosphorylase in Table 25, mean values for the change between 20 and 50ain perfusion in phosphorylase activity (p mol/min/g) were calculated.. Symbols are as in Fig. 25. The line was fitted by regression analysis of values from individual perfusions; r = 0.75, p < 0.001 (22 observations). Bars indicate S.E.M. and the number of perfusions in each group are in Table 25.

OF_

PHO

SPHO

RYLA

SE

163

For Fig. 27 see Over—Leaf

Fig. 27

Relationship be

synthetase and phosphorylase during liver

perfusion. in adrenalectomised rats.

Livers were perfused with glucose plus C3

substrates, with or without pre-treatment in vivo (see Tables

22, 24 & 25). The mean, changes in activities of glycogen

synthetase (% "a" and "a") and phosphorylase between 20 and

50min were calculated (rol/min/g see Fig. 25 and 26). Symbols

are as in Fig. 25. The lines were fitted by regression analysis

of values from individual perfusions, taking into account the

variation in both axes : (a) r = 0.77, p < 0.001 (22 obser-

vations) ;(b) r = 0.75, p< 0.001 (22 observations).

CHANGE IN % GLYCOGEN SYNTHETASE "a"

IN 30 MIN PERFUSION

0 Cn

0

CHANGE IN GLYCOGEN SYNTHETASE "a"

ACTIVITY IN 30 MIN PERFUSION

0

NO

ISIld

213c1

CH

AN

GE IN

PH

OSPH

ORY

LASE A

CTIVITY

C • •

165

For Fig. 28 see Over—Leaf

Fig. 28. • Changes in combined response of both glycogen

synthetase and phoshorla

net glycogen accumulation in the perfused

liver from adrenalectomised rats.

All details and symbols are given in Tables 22,

24 and 25, and Fig. 25. Values for the ratio % glycogen

synthetase "a" / phosphorylase, glycogen synthetase "a" /

phosphorylase were calculated for 20 and 50min liver samples,

and the differences between these values obtained. Bars

indicate S.E.M. and the number of perfusions in each group

are in Tables 25 and 26. Lines were fitted by regression

analysis of values from individual perfusions (a) r = 0.71,

p < 0.001 (22 observations); (b) r = 0.70,p < 0.001 (22

observations).

0 CHANGE IN ENZYME ACTIVITY IN 30 MIN_ PERFUSION

Increase in ( synthetase "a")/(phusphorylase)

2 co

I 1 I I I I I • I 06

Increase in ( o synthetase'crY(phosphorylase)

w.

Uol

iorI

LWID

DIO

O 1-0 O AD O 1.3

O

0

O

167

3.6 AMINO ACID BALANCE IN THE PERFUSED LIVER

1. Amino acid metabolism in fed, starved

and diabetic rat livers.

2. Urea formation in the perfused liver.

3. The role of the liver in amino acid

utilisation.

168

3.6 AMINO ACID BALANCE IN THE PERFUSED LIVER

Net rates of glycogen accumulation were obtained in the

perfused liver of starved rats at rates which occur in vivo (see

Section 3.1). In these experiments synthesis was studied over a

relatively short time period, in the absence of any evaluation of

protein status. Since many hormones are known to have effects on

protein synthesis (enzymes for example) it seemed appropriate that

conditions of protein balance be established in the perfusion so that

any relevance to the action of hormones could be evaluated. One phase

of this study was to assess amino acid release and uptake, at different

initial amino acid concentrations, in different nutritional and

hormonal states. These measurements are relevant to the understanding

of events in livers perfused with amino acids ("supplementation" :

Section 3.4.1), in which net rates of glycogen synthesis were measured,

and so are presented here.

3.6.1 Amino acid metabolism in fed, starved and diabetic

rat livers.

Changes in perfusate amino acids at four times "normal" concentrations

Livers from fed, 48h-starved or streptozotocin -diabetic rats

were perfused in the presence of 30mM glucose, gluconeogenic subStrates

(lactate, glycerol and pymuvate : infused) and amino acids (see Methods

Section 2.3.2). When fed rat livers were perfused in the presence of

amino acids at four times "physiological" concentrations, most were

taken up by the liver (for representative graphs see Fig. 29). Although

there was a delay in the onset of tyrosine uptake (probably due to its,

formation from phenylalanine), after 3h perfusion it reached an

equilibrium at an approximately normal plasma level (Fig. 29). The

0.6

0.3

Cl -U <{

'0 . Z 03 - .

=E <{

=E :::>

0.1 o· w =E

0.9

Fig. 29

TYR

-0 ~~

~00 .~

MET

I LE

~O:::-!b~ o~

°"'-0 ________ 0

ARG

GLU

20 60 120 180

TIME(MIN)

PHE

0.3

0.1

HIS

~

ALA. "/

-11. 3

0--

Metabolism of amino acids during perfusion at high

initial concentrations.

Livers of rats (180-200g; average liver weight about 7g)

were perfused as described in the text vTi th glucose plus a mixture

of lactate, glycerol and pyruvate. Initial amino acid concentration?

vTere about four times "normal" arterial. blood levels. Resul ts are

means of selected values from ~hree perfusions. . Livers vlere from

fed diabetic ( l1 ), 48h- starved normal (0) or fed nomal ( 0 ) rats.

169

170

"branched-chain" amino acids showed little change (for isoleucine,

see Fig. 29), and the amino acids, alanine and glutamic acid, were

transported out of the liver (Fig. 29).

In perfusions of livers from starved or diabetic rats,

the rates of uptake of the amino acids were in general faster than those

,for fed rats, particularly in the case of tyrdsine and lysine (Fig. 29).

Alanine was utilised by the starved rat liver in contrast to those

from fed or diabetic rats (Fig. 29); glutamate was released in both

groups. Glucose and ketone bodies were likely to be major products of the

amino acids used.

The uptake or release of "branched-chain" amino acids by

the liver provides a measure of protein change since they are not

significantly synthesised or utilised by other processes. In these

perfusions the initial concentrations were (approximate nM) : valine,

1200; isoleucine, 400 : leucine, 800. The concentrations in perfused

livers of fed or starved rats did not change significantly during 100min

perfusion whereas the levels increased by 10-20% during perfusion of

diabetic rat livers, suggesting that some proteolysis was occurring.

Release of amino acids during perfusion.

As was seen above, most amino acids (when present at four.

times "normal" concentrations) were taken up by the liver. It was

therefore of interest to evaluate what occurred when no amino acids

were present. In the case of livers from fed animals, most amino

acids normally found in blood appeared in the perfusate and reached

"normal" plasma levels (Table 26). Both glutamic acid and alanine were

however, released in substantial quantities and after 80-100min perfusion

the concentrations were double those of "normal" plasma (Table 26).

171

Table 26. The concentration of amine acids after perfusion

in the presence of "normal" levels or no added

amino acids

Livers were perfused as described in Fig. 29, or

text. Results are means of least 3 perfusions + S.D, except for results

given'for starved or diabetic rat liver perfusions which are means of

2 perfusions. Concentration of amino acids in plasma (1)

No added Amino acids initially "normal" amino acids

Fed Starved Diabetic Fed

Amino acids Final Initial Final

(80-100min.) (5 min) (80-100min)

ASP 69 + 6 68 21 + 4 <20 <20

ASN 44 + 15 130 95 + 20 - _

THRE 103 ± 18 181 62 ± 15 - -

SER 283 ± 20 350 110 + 25 - -

GLU .350 + 52 142 1110 + 156 140 <20

GLN 342 + 32 620 693 + 120 -

GLY . 140 + 34 55o 160 + 25 25 <20

ALA 692 + 82 374 1280 + 54 335 205

VAL 243 + 39 300 279 + 58 260 425

ILE 171 ± 25 125 100 + 25 85 185

LEU 260 14- 52. 266 275 + 30 200 295

MET <20 120 23 + 5 <20 <20

TYR 62 + 15 105 60 + 15 20 <20

PTIE 93 + 12 96 7o + 25 3o 45 HIS 87 + 16 85 65 + 12 20 '35

LYS 220 + 60 420 220 + 40 90 140

ARG <20 110 420 420 420

ORNITH 90 + 20 - - - -

NH3 841 + 75 - - - -

172

The "branched-chain" amino acids were also released, reaching

"physiological" levels, indicating that lysis of endogenous protein

was a major source of all the amino acids released (Table 26).

Livers from starved or diabetic rats released no significant

quantities of amino acids except for valine, leucine and isoleucine

which attained concentrations (results not shown) similar to those

found for the fed liver. Therefore, hepatic proteolysis, under these

conditions, was not altered by starvation or diabetes.

Chan es in erfusate amino acids at "normal" concentrations.

In order to fully assess protein balance in the perfusion,

amino acids were added to the perfusate medium, at "normal" plasma

concentrations. In the perfusions with fed rat livers most amino acids

were taken up (Table 26). Histidine, phenylalanine and glutamine did

not however, change significantly(Table 26). As was noted when four

times "normal" levels of amino acids were added to the perfusate, the

concentrations of the "branched-chain" amino acids, valine, leucine

and isoleucine remained unchanged and alanine and glutamic acid were

released by the liver (Table 26).

When the livers of starved rats were perfused, the amino

acid profiles were similar to those obtained with fed rat livers (Table

26). In general, there appeared to be greater uptake or less release of

amino acids e.g., the concentrations of alanine and glutamic acid did not

change significantly, whereas they were released in fed perfusions

(Table 26). The levels of "branched-chain" amino acids hardly changed.

All amino acids were removed by the diabetic rat liver except

for the "branched-chain" and alanine, which did not alter significantly

(Table 26).

173

3.6.2 Urea formation in the perfused liver

The urea concentrations were measured in the perfusate

plasma, to permit an assessment of nitrogen balance during perfusion.

In perfusions with no added amino acids, the urea level in perfusate

reached 1.5 - 2.4MM (Table 27). In the presence of "normal" levels of

amino acids the urea increased during perfusion of starved or diabetic

rat livers whereas it did not in fed perfusions (Table 27). When four

times "normal" concentrations were in the perfusate, urea increased in

all groups of perfusions, although the urea produced by starved or

diabetic rat livers reached higher levels (Table 27). In all

perfusions with four times "physiological" Amino acids there was a

rapid efflux of urea (Fig. 30), which stabilised during perfusion of

livers from fed rats, but continued to rise during 2h perfusions of

starved or diabetic rat livers.

The urea produced exceeded amino acid uptake (mols N)

in most perfusions (Table 28), except those in which there were four

times "normal" levels of amino acids where nitrogen balance was

attained (Table 28). In perfusions of liver from diabetic rats the

change in amino acid nitrogen exactly corresponded to the production of

urea nitrogen, whereas in those from fed or starved animals the

uptake of amino acids exceeded urea production (Table 28). This latter

finding could reflect the occurrence of hepatic protein synthesis.

174

Table 27. Effect of added amino acids on urea production

in the perfused liver.

Livers were perfused as described in Fig. 29

and in the text. Urea was measured after 80-100min perfusion. Results

are means ± S.E.M. with the number of observations in parenthesis.

EMLIEPLM112.121)

Initial amino acid concentration

Rat None added "Normal" 4 x "Norman'

Starved 1.5 ± 0.2 (3) 3.3 (1) 5.5 + 0.6 (3)

Fed. 1.9 (2) 1.8 (1) 3.7 ± 0.5 (4)

Diabetic 2.4 (2) 3.8 (2) 6.1 + 0.5 (3)

E LU

To

T

A

1 150 50

TIME ( MIN ) 100

Fig. 30. The effect of four times "normal" concentrations

of amino acids on urea production in the erfused

liver.

Livers ( ❑ , fed; 0,48h- starved and A diabetic)

were perfused as described in Fig. 29 and in the text. Samples

of perfusate were removed at the times indicated and assayed for

urea. Results are means of at least 3 observations and the bars

indicate S.E.M.

175

176

Table 28. Nitrogen balance in the perfused liver.

Amino acid change was calculated from measured

initial values and those after 80-100min perfusion; some of the

relevant changes are shown in Fig. 29 and Table 26. Glutamine and .

lysine were considered to provide two mols of nitrogen, histidine

three and arginine four. Urea change was calculated from the

concentration at 80-100min (there being none present initially);

see Table 27.

Unaccounted changes in N during perfusion (which may

reasonably be regarded as reflecting protein metabolism) are expressed

as -ve if amino acid uptake was less than urea production, i.e., if

proteolysis apparently exceeded protein synthesis (and +ve in the

converse case)

Rat

Change in amino- N (m mols)

Amino acid Urea Not accounted for as urea or amino acid.

Amino acids initially four times

Fed

Starved

8.7

14.8

7.4

11.0

+ 1.3

+ 3.8 "normal"

Diabetic 12.2 12.2 + 0.0

Amino acids initially

Fed 0.2 3.6 - 3.4

"normal" Starved 3.2 6.6 - 3.4

Diabetic 3.3 7.6 - 4.3

3.6.3 The role of the liver in amino acid utilisation

177

The present experiments demonstrate the influence of

different hormonal and nutritional states on the uptake of amino

acids by the liver, in conditions where an ample supply of circulating

carbohydrate is available. Breakdown of hepatic protein was minimal

in all groups of perfusions in the presence of amino acids at one or

four times"normal" concentrations, as shown by the insignificant

increases in "branched-chain" amino acids during perfusion. In the

absence of added amino acids, valine, leucine and isoleucine were

however, released by the liver to reach equilibrium at normal plasma

concentrations (present results; see also Mallette et al., 1969).

The presence of glucose and gluconeogenic substrates therefore

inhibited the proteolysis, (measured by "branched-chain" amino acids

released during perfusion) which occurs in the absence of added

carbohydrate, and in the presence or absence of amino acids (Bloxam,

1971). A marked efflux of "branched-chain" amino acids has also

however, been observed in the presence of added carbohydrate and

absence of amino acids (Schimassek & Gerok, 1965), which is inconsistent

with the present findings. One explanation could be the different

perfusion techniques.

The present results confirm that the perfused liver

can regulate the perfusate concentrations of amino acids by net

movement into or out of the cells according to the perfusate

concentrations (see also Bloxam, 1971). Two amino acids which did not

appear to be regulated by the liver were alanine and glutamic acid

which were consistently released during perfusion of livers from,

fed rats, in the presence or absence of added amino acids (see also

178

Schimassek & Gerok, 1965). Such an efflux did not occur when the

livers were from diabetic or starved rats, other than in the presence

of four times "normal" levels of amino acids. It would appear there-

fore that the release of glutamic acid at least, could be a

consequence of the liver glycogen rather than the carbohydrate in

the perfusate. This conclusion is sustained by the results of Mondon

and Mortimore (1967) who observed a notable glutamate release from

the perfused fed rat liver in the absence of added carbohydrate. The

release of alanine on the other hand, is probably a result of the

circulating carbohydrate substrates. It is likely that the efflux of

both glutamic acid and alanine observed from diabetic and starved

rat livers2in the presence of excess amino acids and gluconeogenic

substrates,was due to transamination of pyruvate and Krebs cycle

intermediates occurring faster than these amino acids could be utilised.

The observation however, that alanine was released by perfused livers

from diabetic (but not starved) rats isnot explicable on the known

characteristics of the diabetic state and could imply that alanine is

not an important gluconeogenic precursor in this state.

It has been suggested that there is a permeability barrier

to and from the liver cells for glutamate (Hems et al., 1968).. However,

the present results and those of Schimassek and Gerok (1965) and of

Mondon and Mortimore (1967) illustrate that glutamate can be released

by the liver, and that when present at physiological levels,in the

absence of added substrate2 can be rapidly taken up (Bloxam, 1971).

CHAPTER FOUit

DISCUSSION

180

CHAPTER FOUR

DISCUSSION

4.1 HEPATIC GLYCOGEN METABOLISM IN THE NORMAL RAT

1. The circulating precursors of hepatic glycogen .

2. The role of glucokinase in glycogen accumulation;

3. Control of hepatic glycogen synthesis.

4.2 THE ROLE OP INSULIN IN HEPATIC CARBOHYDRATE METABOLISM

1. Insulin and hepatic glycogen metabolism in the normal starved rat.

2. Hepatic glycogen accumulation in the starved diabetic rat.

3. Properties of glycogen synthetase and phosphorylase in the perfused liver of diabetic rats.

4.3 THP, ROLE OF ADRENOCORTICAL STEROIDS IN HEPATIC GLYCOGEN.

METABOLISM

1. Glucocorticoids and hepatic glycogen metabolism in the normal (starved) rat.

2. Hepatic glycogen accumulation in the starved adrenalectomised rat.

3. The characteristics of glycogen synthetase and phosphorylase in the perfused liver of adrenalectomised rats.

4.4 THE ROLE OF THE HORMONES OF TEE POSTERIOR-PITUITAZY.GLAND

IN REPATIC CARBOHYDRATE METABOLISM

1. The posterior-pituitary gland hormones and the metabolism of liver carbohydrate.

2. The mechanism of vasopressin action.

161

4.1 HEPATIC GLYCOGEN METABOLISM IN THE NORMAL RAT

4.1.1 The circulating precursors of hepatic glycogen

The use of an isolated liver preparation in which net

glycogen deposition occurred at physiological rates and in which all

aspects of glycogen metabolism apparently retained the features present

in vivo, has permitted the characterisation of glycogen metabolism in

rats. An initial question concerns the nature of the circulating

precursors of glycogen. The experiments reported (Section 3.1.2)

suggest that much of the hepatic glycogen that accumulates in the rat

on re -feedingl after a period of starvation,could be initially derived

from gluconeogenesis (defined as the new synthesis of glucose in either

mono- or poly saccharide form). This follows from the facts that the

fastest rates of glycogen deposition in the perfused liver were obtained

with gluconeogenic precursors in the perfusion medium as well as glucose,

and that there was no net uptake of glucose in this situation, despite

its high concentration (25-30MM). These characteristics of hepatic

glycogen synthesis may reasonably be presumed to exist in vivo, because

the rate of glycogen deposition, its dependence on glucose concentration,

and time course, were similar in the perfused liver and in the intact

rat.

It is reasonable to suppose that gluconeogenesis could have a

major role in the redeposition of hepatic glycogen after re-feeding

with material other than glucose. It is also possible that the

continuation of gluconeogenesis (from "endogenous" precursors) could

contribute to hepatic glycogen accumulation after the post-starvation

ingestion of glucose. In this situation, the increase in circulating

glucose concentration would relieve the liver of its function to release

free glucose into the blood, and the newly formed hepatic glucose

182

phosphates would be directed to form glycogen. The present experiments

suggest that this could be the initial sequence of events in vivo on

re-feeding with glucose. This possibility was suggested from experiments

on intact rats by Olavarria et al., (1968),* and was also raised by

Jeffcoate and Moody (1969) and Zaragoza -Hermans and Felber (1972).

Although the concentrations of circulating precursors in

the present experiments were high compared with those in the intact rat,

the findings may nevertheless be related to the starvation-re-feeding

situation in vivo. The question of particular interest is whether glucose

in the blood, after ingestion by a starved rat, is taken up by the liver

to form glycogen. In the perfusion experiments, glucose and gluconeogenic

precursors were present at near-saturating concentrations and the liver

exhibited a total preference for the gluconeogenic substrates. In the

intact 48h- starved-refed rat, although the blood concentrations of

gluconeogenic substrates are lower than were employed in perfusion),

so is that of glucose. Hence it is likely that in the initial phase of

re-feeding with glucose in vivo, the preference for gluconeogenic

precursors would prevail and that net glucose uptake by the liver would

be negligible.

During starvation, glucose tolerance is impaired in comparison

with the fed state. The present results suggest that, in the initial

phase of assimilation of a glucose load by tissues of starved animals,

there is no significant net uptake of glucose by the liver, although

there may be a marked diminution in the hepatic release of glucose into

blood, caused largely by the increased concentration of blood glucose

and perhaps by insulin. This conclusion is in accord with the main

explanation for the glucose intolerance of starvation proposed by Mahler

and Szabo (1970).

183

On ingestion after starvationl glucose is degraded by

extrahepatic tissues into potentially gluco genic precursors such

as lactate and alanine. These precursors, diluted in the general pools

of "endogenous" precursors, would eventually become incorporated into

liver glycogen. The extent of this process has been measured by

Friedmann et al., (1965), and was estimated at 20-40% of the glycogen

synthesis. This result is not incompatible with the present conclusion

that hepatic glycogen accumulation, after ingestion of glucose, may

initially derive largely from the continuing flux of "endogenous"

gluconeogenic precursors.

After a period of deprivation of dietary glucose, hepatic

metabolism adapts on eventual re-feeding with glucose, so as to decrease

the rate of gluconeogenesis. The present experiments suggest that

the initial phase of such an adjustment between "starved" and "fed"

states (defined with regard to dietary glucose) need not involve a

cessation of hepatic gluconeogenesis but rather its continuation for

perhaps several hours, contributing to the reaccumulation of glycogen.

During this time the activity of glucokinase in liver may slowly

increase (Salad et al., 1963; Sharma et al., 1963). In the event of

persistent repletion with dietary glucose, net uptake of glucose by the

liver could eventually supervene. Such a slow adaption to re-feeding

with glucose is likely to be of greater value to an animal than a more

rapid response to a possibly transient surge of glucose in the portal vein.

The existence of net gluconeogenesis in livers from fed rats

is difficult to demonstrate, in the presence of the usual associated

glycogen breakdown. The present experiments (Section 3.2.1) with a

glucogenic mixture, in which net glycogen breakdown was suppressed by

high glucose concentrations, showed however, that net gluconeogenesis

can occur in such livers and is not inhibited by high glucose concentrations.

184

The existence of gluconeogenesis in the fed state is in general

agreement with previous experiments with 14C-labelled gluconeogenic

precursors (Clark et al., 1974;EXton et al., 1972b)..

The results obtained with overnight-starved rats (Section

302.2).confirm the importance of gluconeogenesis in glycogen deposition,

although the influence of gluconeogenic precursors on hepatic glycogen

synthesis is much less marked and that of glucose more, than in 48h-

starved rats. In this condition there would not have been time for a

full adaptation of the enzyme3for the metabolism for 03-substrates,

which occurs on longer starvation. Even between 18 and 22h following

food deprivation, there was an increase in the capacity to form glycogen

from the mixture of glycerol, lactate and pyruvate. On the other hand,

the accumulation of glycogen in perfusions containing fructose showed no

difference between 18 and 48h- starved states, in agreement with the

general concept that adaptations in this part of the gluconeogenic

pathway are not as great as in the enzymes which metabolise pyruvate.

On considering the calculated rates of glucose release from

these livers, i.e., fed, 18h- starved and 48h- starved, adaptations to

starvation in the pathway of gluconeogenesis can be seen. The rate of

glucose release from the fed rat liver was 4.0iumol/min which is about

half the rate of gluconeogenesis in matched 48h- starved rats (about

8.0 prnol/min per liver). In overnight starved rats the rate was 5.6)1=1/

min per liver. This difference between fed and starved animals is of

the same order as that obtained when fed rats were depleted of liver

glycogen before liver perfusion (Ross et al., 1967). It would appear

therefore that there is some adaptation of the pathway of gluconeogenesis

to different nutritional states but that the capacity for the process is

always present. This is in keeping with the assayable activities of

regulatory enzymes of gluconeogenesis, and the extent of changes on e.g.,

starvation.

185

In general, the precursors of hepatic glycogen appear to

be gluconeogenic, the net contribution of glucose being minimal perhaps

even in the fed state.

4.1.2 The role of glucokinase in glycogen accumulation

Hepatic glucokinase activity is not always sufficient to

account for the uptake of glucose to form glycogen after the post-

starvation ingestion of glucose. It is known that glucokinase activity

falls on starvation and diabetes.(Salas et al., 1963; Walker & Rao, 1964),

although levels are sufficient to support some phosphorylation of glucose,

they would not allow net glycogen deposition. The experiment reported

14 1 \ with k C) -glucose in the perfusion medium (Section 3.1.2) showed that

glucose uptake (uni -directional, rather than net) did occur during

maximal glycogen synthesis from gluconeogenic precursors in the liver

from 48h- starved rats,and that glucose as a carbon source contributed

about one-third of the total carbon.

Alternative pathways of glucose uptake have been suggested

(see Friedmann et al., 1967; Ryman & Whelan, 1971) to circumvent the

low activity of glucokinase. The present experiments may however, resolve

this difficulty, in suggesting that gluconeogenesis could make a major

contribution to hepatic. glycogen synthesis on administration of glucose to

starved rats or insulin to diabetics. Thus there is no requirement

for glucokinase activity to be as rapid as the rate of glycogen deposition.

If indeed the net uptake of glucose by the liver to form

glycogen is of minor importance, even in fed non-ruminant mammals, the

question arises of the significance of the glucokinase enzyme. This

could reside in the control of glycogen metabolism by glucose, which

could involve phosphorylation of glucose (see Section 4.1.3).

186

4.1.3 Control of hepatic glycogen synthesis

The influence of increasing glucose levels has been

extensively studied in various experimental systems and it has been

found that glycogen synthesis is enhanced in a dose-dependent manner*

(see Introduction, Section 1.3). A similar result, in the presence of

gluconeogenic substrates, is reported here (Section 3.1.5).

On glucose ingestion after starvation the initiation of

glycogen synthesis is due to activation of glycogen synthetase (see

Hers et al., 1970, for review; Hornbrook, 1970) and inhibition of

phosphorylase (Hornbrook, 1970). The present results suggest that this

initiation of glycogen synthesis could largely be a direct consequence

of an increase in glucose concentration in the hepatic portal vein,

since maximal glycogen accumulation required the presence of glucose

in the medium, but no added hormones (Section 3.1.2). The rate of

total gluconeogenesis (glucose plus glycogen) was not significantly

altered by the presence of glucose in the medium (as was observed by

Exton & Park, 1967; Haft, 1967) which is in accord with the possibility

that glucose (or its metabolic products) may primarily activate the

glycogen synthetase system. Such activation of glycogen synthetase

(concomitant with an inhibition of phosphorylase activity) by glucose

(Buschiazzo et al., 197.0; Glinsmann et alo, 1970; Miller et al., 1973)

and fructose (Valli et al., 1974) has been demonstrated in the isolated

perfubed liver. It has also been demonstrated that by increasing the

glucose concentration a more extensive activation of glycogen synthetase

and inhibition of phosphorylase is obtained (Glinsmann et al., 1970).

It appears from the present results that gluconeogenic substrates,

including fructose,enhance the response of glycogen synthetase to glucose,

since there was less activation by glucose or gluconeogenic substrates

alone. The C3-substrates did not however, enhance the response of

187

phosphorylase to glucose (Section 3.1.5).

The "a" and "b" forms of glycogen synthetase and phosphorylase

are interconverted by protein kinases and phosphatases (see Introduction,

Section 1.2) and glucose appears to affect the activity of these glycogen

metabolising enzymes by increasing the activity of glycogen synthetase

phosphatase (DeUulf & Hers, 1967a & 1968b) and phosphorylase phosphatase

(Stalmans et al., 1970 & 1974a). Since glucose action on phosphorylase

phosphatase decreases the amount of phosphorylase "a" (active form of the

enzyme) this in turn will cause deinhibition of glycogen synthetase

phosphatase by phosphorylase "a" (Stalmans et al., 1971 & 1974a).

The question arises as to whether it is glucose specifically,

or a metabolite which initiates glycogen synthesis. The glucose

concentration-dependence of synthesis suggested that glucose phosphorylation

may be relevant in the initiation of synthesis, since it resembled that of

glucokinase (Walker & Rao, 1964). Consideration of the role of glucose in

inducing or initiating processes in other tissues may shed light on this

question. For example, studies on glucose-induced insulin release in the

pancreas may bear on this problem. The effect of glucose on insulin release

is critically dependent on extracellular concentrations of the sugar and it

was therefore suggested (Cerasi & Luft, 1970; Landgraf et al., 1971) that

pancreaticA5-cells possess a glucoreceptor system (which, through molecular

interaction with glucose at an appropriate concentration of glucose; leads

to activation of insulin release). It has also been proposed however, that

it is a glucose metabolite that activates insulin release (rather than glucose

itself). These hypotheses have been termed regulator site and substrate

(or metabolite) site, respectively (see Randle & Hales, 1972 for further

discussion). The initial suggestion for the operation of the substrate site

mechanism came from the observation that sugars readily metabolised by

mammalian tissues (for example

188

glucose, mannose and fructose) elicit insulin release (see MacDonald

et al., 1975). Sugars that are not readily metabolised (e.g.,

galactose and 5-0-methylglucose) or that are only phosphorylated

(2-deoxyglucose) did not stimulate insulin release (Coore & Randle,

1964; Grodsky et al., 1965). The regulator (glucoreceptor) site

hypothesis has obvious attractions, for it involves direct interaction

of glucose with a specific receptor. If such a receptor exists, it

would have a pattern of specificity however, which has not been found

among glucose transporting systems or enzymes. At present therefore,

evidence suggests that it is a metabolite of glucose (probably not glucose

6-phosphate), which activates insulin release.

As has been seen in the present work and noted above, some

phosphorylation of glucose occurs during glycogen synthesis. Since

the rate of glycogen accumulation in the perfused liver was dependent

on the glucose concentration over the range 10-30mM, which corresponds

to the !Km? (for glucose) of hepatic glucokinase (Di Pietro et al.,

1962; Walker & Rao, 1964), phosphorylation of glucose and perhaps

further metabolism of the hexose (as was proposed above with regard to

the stimulation of insulin secretion) may be implicated in the initiation

of hepatic glycogen synthesis by glucose. This would accord a role to

glucokinase in the regulation of glycogen metabolism. A number of

glucose-like substances (those readily metabolised, non-readily

metabolised or only phosphorylated) do not, however, substitute for

glucose in shortening the lag phase of glycogen synthetase activation in

vitro (DeWulf et al., 1970) or inactivate glycogen phosphorylase (Stalmans

et al., 1970). It has recently been shown that a few derivatives of

glucose do, however, stimulate the phosphorylase phosphatase reaction,

1, 5 -anhydroglucitol being the most effective (Stalmans et al., 1974b).

This indicates that it maybe glucose itself and not a metabolite, which

189

is involved in the activation of glycogen synthetase, inhibition of

phosphorylase and hence initiation of synthesis.

The concentration of glucose 6-phosphate in the cell has •

been frequently considered as an important factor in the control of

glycogen synthesis from UDPG. According to this hypothesis, the leyel

of glucose 6-phosphate in the liver would be increased after a glucose

load, as a direct result of hyperglycaemia, whereas the concentration of

UDPG would possibly be lowered. However, both metabolites decrease in

the liver of fed animals and the concentration of UDPG is diminished

whereas that of glucose 6-phosphate is already low, in the fasted mouse

(DeWulf & Hers, 1967a). These observations rule out glucose 6-phosphate

as a stimulatory agent. In vitro studies have also mitigated against

the importance of glucose 6-phosphate as an activator of glycogen

synthetase "a" and "b" and indicate that Pi is the more important enzyme

regulator (DeWulf et al., 1968). The fact that both UDPG and glucose

6-phosphate decrease after a glucose load indicate that a pull mechanism

exists such as would be obtained by a stimulation of the last steps of

glycogen synthesis. In the present work a decrease in the hepatic UDPG

concentration was observed when glycogen synthesis was maximal, although

there was an increase in glucose 6-phosphate (Section 3.1.5).This latter

finding is probably due'to the presence of gluconeogenic substrates in

the perfusion medium and suggests that in these conditions glucose 6-

phosphate could affect the activity of glycogen synthetase. For

example, the "b" form could be activated by glucose 6-phosphate, since

conversion to "a" is never complete (up to about 80% in the present

experiments).

190

It is not known how glucose directs hexose phosphates

to glycogen rather than glucose. It is known however, that a "pull"

on glycogen precursors would occur due to glucose-activated glycogen

synthetase, and it is possible that the inhibition of glucose 6-

phosphatase is involved (see Cahill et al., 1959; Ryman & Whelan, 1971).

It was found in the present work that when glycogen accumulation

occurred during perfusion there was less glucose release, but that the

total glucose formed (glucose and glycogen) at different initial glucose

concentrations, was constant (except when the glucose concentration was

40MM or there were no gluconeogenic precursors). This could indicate

that the perfusate glucose has to reach a certain concentration before

. the hexose phosphates are directed to glycogen in preference to glucose.

It is noteworthy that high concentrations of circulating glucose (40-50MK)

inhibited glycogen deposition although not gluconeogenesis : this could

.reflect an osmotic action of such a high solute concentration, or more

complex effects of glacose.Overall several aspects of the regulation of

hepatic metabolism by glucose remain unsolved (as in the case of

initiation of insulin secretion by glucose in the pancreatic /3 -cell)

191

4.2 THE ROLE OF IlibULIN IN HEPATIC CARBOHYDRATE METABOLISM

402.1 Insulin and hepatic glycogen metabolism in the •

normal starved rat.

There is controversy about the possible existence of a rapid,

direct action of insulin on the liver in post-starvation glycogen

deposition. In support of this possibility, hepatic glycogen synthetase

can be activated by insulin in certain conditions (Bishop et al., 1971),

glycogen synthesis can be prevented by anti-insulin serum (Steiner, 1964)

and insulin can stimulate glycogen synthesis from glucose in the isolated

perfused liver of starved rats (Sakai et al., 1958). It is established

that insulin can inhibit hepatic glycogen breakdown in the perfused

liver in some conditions (see Mondon & Burton, 1971, for references), as

it can in the intact animal (see Madison (1969) and Steele (1966)for

reviews). However, there is also evidence against a direct hepatic

action of insulin in the initial phase of post-starvation glucose

assimilation. For example, insulin does not activate glycogen synthetase

in the perfused rat liver (Glinsmann et al., 1970), and anti-insulin

serum does not alter the immediate hepatic fate of administered glucose

in rats (Moody et al., 1970). In rats with portal-caval anastomosis,a

combination of high circulating insulin an normal glucose concentrations

is associated with decreased hepatic glycogen synthesis (Assal et al.,

1971).

Since the immediate post-starvation synthesis of glycogen

appears to be largely a result of gluconeogenesis (even after glucose

ingestion), insulin, which can inhibit gluconeogenesis in the isolated

liver (Exton et al., 1970), would not be expected to stimulate this

process. In the present experiments with the perfused liver, glycogen

192

synthesis was rapid in the absence of added hormones, and insulin did

not influence glycogen synthesis under optimum conditions (Section 3.1.3).

Hence in the immediate post-starvation situation it appears unlikely

that circulating insulin exerts a major direct stimulatory effect on

liver glycogen synthesis. This is in general agreement with the

conclusions of Glinsmann et al., (1970), Hers et al., (1970) and

Moody et al., (1970). However, it is possible that insulin may exert

a moderate hepatic effect during post-starvation glycogen deposition,

in view of its known action in preventing glycogen breakdown. In the

present experiments insulin( albeit at a high concentration) produced

a moderate stimulation of glycogen deposition in suboptimal conditions,

i.e., low glucose concentration or no C3-substrates. Also, the

present data do not exclude a possible hepatic action of insulin in vivo

in combination with other hormones (e.g., in counteracting the glycogenolytic

action of glucagon Glinsmann & Mortimore, 1968; Mackrell & Sokal, 1969).

Regarding the effect of anti-insulin serum, obtained by Steiner (1964),

this could reflect extrahepatic events that indirectly resulted in a

glycogenolytic action on the liver, or the relatively long duration of

the experiment (3.5h), since the present results are not incompatible

with a longer-term action of insulin on liver glycogen metabolism. Evidence

discussed in the next section supports the first of these possibilities.

193

4.2.2 IleaaLticalysogen accumulation in the starved diabetic rat.

Imaientofheat3.c12 bhesisindiabetesrm . For

a variety of reasons (see Introduction, Section 1.4.1) it has not been

clear whether there is a significant and inherent alteration in

maximal rates of net hepatic glycogen accumulation in diabetes. The

streptozotocin-diabetic animal provides a suitable model for the

clarification of this problem (Junod et al., 1969). The changes in

glycogen metabolism described previously (Section 3.4) reflect the

consequences of insulin-deficiency diabetes, rather than hepatic toxicity

of streptozotocin since they were reversed by insulin and hexoses in

vivo.

The results (Section 3.4.1) show that there is a marked

inherent impairment in the maximum rate of net glycogen accumulation

in the liver of starved diabetic animals. In all groups of perfusions

of the livers of diabetic rats (in the absence of pre-treatment in vivo),

rates of net glycogen synthesis were low compared to those in normal

(starved) rats. This result confirms, by the measurement of net glycogen

deposition in optimal conditions, the inherent impairment of hepatic

glycogen synthesis in diabetes which has been inferred from measurements

of incorporation of 14C from 14

C- labelled precursors into glycogen in

the perfused liver (Exton et al., 1972b & 1973a) or in liver slices (for

reviews see Levine & Fritz, 1956;.Renold et al., 1956; Steiner, 1966).

In general, measurements of net glycogen synthesis in intact

diabetic rats (Friedmann et al., 1963 & 1967; Hornbrook, 1970; Longley

et al., 1957), do not show such a marked impairment of synthesis as that

revealed in the perfused liver. Thus the present perfusion experiments

demonstrate the extent of the potential decrease in the maximal rate of

glycogen synthesis in diabetes, that is inherent to the liver, and its

194

rapid reversibility (by treatment in vivo).

In experiments with intact diabetic animals, the decrease

in the rate of hepatic glycogen synthesis can be shown by measurements

of incorporation of 14C from C from C- labelled precursors into glycogen

(Friedmann et al., 1970; for reviews of earlier work, see Renold et

al., 1956; Steiner, 1966). The general implication of these

observations is that in diabetes, the hexose -phosphate products of

gluconeogenesis (produced at an increased rate) are directed towards

free glucose formation rather than glycogen (although the proportional

contribution of gluconeogenesis to glycogen formation is increased :

Stetten & Boxer, 1944). Results obtained in the perfused liver confirm

this suggestion, in that during maximal glycogen synthesis, the net carbon

sources of glycogen are gluconeogenic precursors, in either normal

(starved) rat livers (see Section 3.1.2) or in diabetic rat livers

(see Section 5.4.1). Net glucose uptake by the liver is not of significance

in diabetes. However, glucose must be present during perfusion to permit

glycogen deposition, even if net uptake of glucose does not occur. The

mechanisms underlying this requirement for glucose are not clear (see

also Seglen, 1974).

Studies of net glycogen metabolism in the liver of intact

diabetic animals show that as well as there being a defect in the rate

of glycogen accumulation (see Results, Section 3.4.4), there is a

limitation in the amount of glycogen which can be stored (Friedmann,

et al., 1967; Hornbrook, 1970). The present experiments did not however,

bear on this latter aspect of hepatic glycogen metabolism in diabetes.

Restoration of hepatic :1 cocren accumulation in diabetes.

Restoration of net glycogen synthesis in the perfused liver of

starved diabetic rats was achieved by two procedures in vivo : (i) pre-

' treatment with glucose plus fructose, (ii) insulin pretreatment (50-75min).

This effect of insulin would be expected, as it was repeatedly been shown

195

that the consequences of diabetes due to pancreatic A -cell toxins may

be reversed by insulin in vivo.

The results shed light on the origin of the inherent

alteration in hepatic glycogen synthesis rates in diabetes. Since

insulin in vitro produced no restoration of synthesis in perfusions

lasting up to 100min , but induced restoration within about 75min ,

if administered in vivo, it follows that a decline in the short-term

direct actions of insulin on the liver cannot be a major contributory

factor (Levine & Fritz, 1956). This conclusion is in agreement with the -

lack of action of insulin under optimal conditions for glycogen synthesis

in the perfused liver of normal (starved) rats (see Discussion, Section

4.2.1) and is not vitiated by the observation that insulin can suppress

the breakdown of glycogen in the perfused liver (see Exton et al., 1970)

or in hepatocytes (Akpan et al., 1974). Similarly in studies with 140-

labelled precursors, impairments in glycogen synthesis were not corrected

by insulin in the perfused liver (Exton et al., 1973a; Haft, 1968) or

liver slices (Renold et al., 1955).

Although the rats employed in the present experiments were

diabetic (based on the normal criteria), there was no necessity for

insulin maintainence, and thus they were not severely diabetic. However,

the impairment in hepatic glycogen synthesis was marked (albeit rapidly

reversible). The liver may therefore be sensitive (indirectly) to small

falls in plasma insulin (in the intact animal). It is likely that

starvation of the rats, in further lowering the circulating insulin level,

contributed to the development of the alteration in glycogen metabolism.

Thus it might be expected that the consequences of diabetes (insulin lack)

on glycogen metabolism in "fed" rats may be less consistent. These

factors might explain some of the. variability in resported effects of

insulin deficiency on hepatic glycogen metabolism.

J 196

The present experiments also show that hepatic glycogen

synthesis is not sensitive to a decline in direct long-term insulin action.

This follows because the alterations in diabetic animals were reversed

within 75min by procedures in vivo (tantamount to re-feeding), through

mechanisms which did not include direct hepatic insulin action.

Among possible factors acting in vivo, by which insulin may

affect the liver indirectly, the present experiments exclude any simple

role for adrenal corticosteroids (since hydrocortisone did not restore

synthesis of glycogen), or a fall in the blood concentration of glucagon

or free fatty acids, since prolonged perfusion in the absence of these

agents did not result in increased net glycogen accumulation. Hence

extra-hepatic effects or unknown co-factors appear to be implicated in

those rapid insulin actions which ultimately affect liver glycogen

synthesis in intact animals. These mechanisms are entirely obscure.

The restorative effect of hexoses on net glycogen synthesis

(measured during eventual perfusion) appears to involve a specific action -

of fructose. Thus the action of fructose plus glucose could not be

reproduced by an equivalent quantity of glucose, and anti-insulin serum

did not completely prevent their action. Also, in the intact diabetic

rat,glucose alone or plus a mixture of C3 -substrates, did not restore net

glycogen synthesis as effectively as did glucose plus fructose. This

effect of fructose is reminiscent of its action in diabetic rats in

restoring rates of fatty acid synthesis (Baker et al., 1952). The

mechanism of its action is likely'to include production of glucose (and

related metabolites) within the liver, since the liver is a main site of

fructose utilisation, and in diabetes this process is not impaired in the

same way as that of glucose assimilation (Chernick & Chaikoff, 1951;

Miller et al., 1952). Additional factors may be implicated in the

restorative effect in vivo of fructoseand glucose. One such factor is

197

likely to be insulin, which was effective over the same period

in vivo, and the restorative action of the hexoses was shown to be

partially prevented by anti-insulin serum. Insulin secretion, although

impaired, can be stimulated by administration of glucose to streptozotocin

-diabetic rats (Junod et al., 1969 )and it is known that fructose

may potentiate this action (Curry et al., 1972; Dunnigan & Ford, 1975).

4.2.3 Properties of almogmEalletase and phosphorylase

in the erfused liver of diabetic rats.

There could be various origins for the impairment in the

capacity for net glycogen synthesis in the liver of diabetic rats. The

present experiments suggest that the explanation may reside at least

partly in an inadequate responsiveness of the hepatic glycogen synthetase

system to substrates. Thus during perfusion of liars from diabetic rats,

in the absence of pre-treatment, there was an impairment in the increase

of glycogen synthetase "a" (active form of the enzyme : Herb et al.,

1970) which was observed in response to glucose plus 03-substrates in

normal rat livers. Similarly,if glucose alone is added to the medium

perfusing the liver of diabetic. rats (i.e., in conditions, which are not

conducive to maximal glycogen synthesis) impairment in the response of

synthetase and phosphorylase to glucose may be observed (Miller et al.,

1973). In the present experiments, the impairment in response of

synthetase to substrates in diabetes was not complete, since fructose (in

the presence of glucose) was able to activate synthetase in the perfused

liver of diabetic rats (in the absence of pre-treatment in vivo).

An impaired activation of glycogen synthetase was also demonstrated in

the intact rat, being most noticeable in the relative lack of response to

glucose plus fructose, compared to that in normal (starved) rats. This

is in general agreement with the results of Steiner (1964 & 1966),

Kreutner & Goldberg (1967) and Miller et al., (1973). The response of

198

phosphorylase to glucose and fructose was not impaired.

Insulin administration in vivo rapidly restored the sensitivity

of glycogen synthetase and phosphorylase to glucose plus C3-substrates

in the diabetic liver (as it restored the associated low rates of net

glycogen synthesis). This is unlikely to reflect a direct hepatic action

of insulin, as was shown above (Section 4.2.2); thus the high proportions

of synthetase "a" in the livers perfused with supplemented medium

(including insulin) probably involved the action of fructose, observed

in perfusions with fructose and glucose alone. Insulin effects on

glycogen synthetase in isolated liver preparations (in contrast to frog

liver : Blatt & Kim, 1971a) are in general not extensive (Akpan et al.,

1974; Miller & Larner, 1973). An exception to this generalisation is

that insulin can counteract the rapid (inhibitory) actions of glucagon or

adrenalin in the perfused liver on glycogen synthetase (Hostmark, 1973;

Miller & Lamer, 1973); this type of action may have a role in the rapid

effect of insulin in intact diabetic animals, in which circulating glucagon

concentrations are increased (Unger, 1972).

If direct, short-term insulin action on hepatic glycogen synthesis

(and synthetase)is not significant (in rats at least) then it follows that

the effect of insulin on hepatic glycogen synthetase in intact animals

(Bishop et al., 1971; Blatt & Kim, 1971b; Miller & Lamer, 1973; Nichols

& Goldberg, 1972) does not involve direct action. This conclusion endorses

in general that derived by Hers et al., (1970) from experiments in intact

mice. Similar considerations apply to the decrease in hepatic synthetase

phosphatase activity of diabetic animals (Bishop, 1970; Gold, 1970b);

this activity can be restored by insulin in vivo, but perhaps not as a

result of direct insulin action on the liver.

199

The behaviour of glycogen synthetase and phosphorylase in vivo

and in perfusion suggest that fructose can exert specific effects on

their activities (see Van delBerghe et al., 1973) and in restoring their

efficacy in the liver of diabetic rats. Thus fructose in combination

with glucose in the perfusate produced the highest proportional

activities of glycogen synthetase "a", in the liver of normal rats (in

vivo or in perfusion). Also, in diabetic rats, fructose plus glucose

in vivo resulted in the highest initial glycogen synthetase "a" in

subsequent perfusiohs containing fructose. Anti-insulin serum did not

completely prevent the action of fructose and glucose in vivo in

increasing the response of synthetase "a", during perfusion with C3-substrates,

suggesting that the effect of fructose (plus glucose) in correcting the

impaired sensitivity of synthetase and phosphorylase to substrates in

diabetes involves actions in vivo additional to the stimulation of insulin

secretion. These are likely to include specific hepatic effects, such

as a decline in the hepatic cyclic AMP content (which is raised in diabetes :

Pilkis et al., 1974) and an increase in hepatic fructose 1-phosphate contest;

both these changes occur in response to fructose, being relevant to the

associated decrease in glycogen phosphorylase activity (Thurston et al.,

1974; Van den Berghe et al., 1973).

The attainment of a.high proportion of synthetase "a" during perfusion

with fructose shows that the enzymic apparatus for glycogen synthesis is

not fundamentally lacking in the diabetic rat liver (see also Hornbrook,

1970). This also follows from the rapidity of the correction of impaired

glycogen synthesis by treatment in vivo. Yet, such activation by fructose

was not a sufficient condition for maximal net glycogen accumulation. Thus

in diabetes, the close relationship between synthetase "a" and the rate

of hepatic glycogen synthesis need not hold. A possible explanation for

this is the following. Maximal glycogen synthesis in the livers of normal

200

(starved) animals involves glucose and other carbon sources acting in

combination. Glucose, while not providing carbon for glycogen synthesis,

appears to exert an "initiatory" effect on synthesis by influencing the

fate of newly-synthesised hexose phosphate, i.e., in directing this

hexose to glycogen rather than free glucose. Presumably, in the diabetic

rat liver, a component of this overall process is impaired. This

deficiency cannot be in the process of gluconeogenesis, which is enhanced

in diabetes. Also, the effect of fructose in the perfused liver suggests

that the glycogen synthetase system is not defective. The defect which could not

be corrected in the perfused liver may therefore involve the initiatory or

cofactor role of,glucose, which is implicated in net glycogen deposition

in the normal liver and apparently in maintaining the full sensitivity of

synthetase (see Section 4.1.3). This would be in accord with the

established impairment in hepatic glucose uptake in diabetes (see Renold

et al., 1956; Steiner, 1966), and would also explain Why the hepatic

glycogen deposition in diabetes can be corrected more rapidly than is

accounted for by the restoration of glucokinase activity (Steiner, 1964)

since this regulatory role of glucose could involve free glucose (Hers

et al., 1970) as well as metabolic products. The nature of the glucose

effect on the liver is obscure; correction of this impaired glucose

effect in diabetes also involves factors which are not clear, including

insulin, but not solely (if at all) as a result of direct hepatic insulin

action.

201 •

4.3 TBE ROLE OF A METABOLISM

AL CORTICAL STEROIDS IN HEPATIC GLYCOGEN

4.3.1 Glucocorticoids and hepatic glycogen metabolism in the normal starved rat.

The stimulation of hepatic glycogen synthesis by glucocorticoids

was described in 1940 by Long, Katzin and Fry. Since then it has been hydro

shown that/col*;scale increases the conversion of circulating glucose into

liver glycogen (Ashmore et al., 1961; Gold & Segal, 1966), by activating

glycogen synthetase (DeUulf & Hers, 1967b, and 1968b) and inhibiting

phosphorylase (Stalmans et al., 1970).

In the present experiments with the perfused liver from

starved rats, glycogen synthesis was rapid in the absence of added hormones

and hydrocortisone did not alter synthesis under optimal conditions (see

Fig. 21, Section 3.4.1). This result is not however, incompatible with

glucocorticoids having a role in glycogen synthesis in the longer-term or

in vitro in suboptimal conditions, but suggests, in particular, that they

have no direct short-term effect on hepatic glycogeneds in normal rats.

4.3.2- Hepatic glYcogen accumulation in the starved

and adrenalectomised rat.

Impairment of hepgaticicoens -yadrenano.

Aspects of the role of adrenal cortical hormones in glycogen metabolism

are not fully clear. For example,'it is uncertain whether alterations

observed in hepatic metabolism,after adrenalectomy, are inherent to the .

liver.

The present results with starved adrenalectomised rats showed

that there was a marked inherent impairment in glycogen synthesis in all

groups of liver perfusions, even after pre-treatment in vivo with

hydrocortisone and or insulin, for periods of up to lh. Since the

202

metabolic changes due to adrenalectomy are not rapidly corrected (by

substrates in vivo, or even by the replacement of the missing hormone),

this hormone-deficient state would appear to be a more severe condition

than diabetes, where "normal" rates of glycogenesis were obtained after

lh treatment in vivo with glucose and fructose (see Discussion, Section

402. ).

This total loss of glycogen synthetic capacity represents a

much more severe alteration in hepatic metabolism following adrenalectomy

(plus starvation) than has been reported so far for any other metabolic

process (e.g., gluconeogenesis, ketogenesis, fatty acid metabolism) . The

impairment in gluconeogenesis,for example,which is relevant to glycogen

deposition in the present conditions (see Discussion, Section 4.1.) is

not severe,and is insufficient to explain the total loss of glycogen

synthetic capacity after adrenalectomy.

Hydrocortisone,in vitro, did not increase glycogen accumulation

in short or longer-term experiments,confirming the lack of a direct hepatic

effect on this process (see also Exton et al., 1973b). A similar lack of

response of glycogen synthesis to hydrocortisone was observed when the

livers of starved or diabetic rats were perfused. Although steroid-induced

glycogenesis has been postulated to be mediated by insulin (Exton et al.,

1973b, Hornbrook, 1970; Kreutner & Goldberg, 1967; Nichols & Goldberg,

1972), insulin did not have any effect when added in vitro. The nature of

the insulin effect in the intact animal is therefore obscure.

The marked impairment in glycogen accumulation observed in

perfusion was confirmed in vivo, showing that the impairMent was not a

phenomenon of perfusion as an experimental system or due to the omission in

of circulating factors. A similar impairment/in vivo glycogen synthesis

(although not glucose synthesis) from 140-labelled alanine or pyruvate,

has been observed (Friedmann et al., 1965). There is evidence however,

to suggest that glycogen synthesis is rapidly restored in the intact rat, for

203

after intragastric glucose, hepatic glycogen levels increase, although

not until 2-3h. Such "normal" rates of glycogenesis may fall off after

8h (Friedmann et al., 1967). It was therefore suggested that the defect

in adrenalectomy resides in the storage of glycogen rather than in its

synthesis.

Restoration of hepatic glycogen synthesis in adrenalectomy.

Restoration of net rates of hepatic glycogen accumulation in perfusion

were achieved by a number of procedures in vivo (a) pre-treatment with

hydrocortisone for 3-4h; (b) pre-treatment with hydrocortisone, fructose

and glucose for 2h or (c) pre-treatment with insulin, fructose and glucose

for 2h.

(a) The role of glucocorticoids in hepatic glycogen synthesis.

The restorative effect of hydrocortisone on hepatic glycogenesis was as

previously reported (Kreutner & Goldberg, 1967; Nichols & Goldberg, 1972).

However, since the hormone in vitro did not affect synthesis in perfusions

lasting up to 4h (treatment in vivo for this time restoring synthesis), /apparently

the restorative effect cannotbe due to a direct action on the liver (see

above).

time course of The/restoration of net rates of glycogen synthesis by steroid

was shortened by the addition of fructose and glucose during pre-treatment.

These hexoses were found to restore synthesis in the livers from diabetic

animals and it was suggested that fructose may have a specific action,

which could include stimulation of insulin secretion (see Discussion, Section

4.2. ). In adrenalectomised rats, insulin secretion is impaired (Malaisse

et al., 1967), the response of the pancreas to glucose being restored by

hydrocortisone. Since the adrenalectomised rats used in them studies

were starved for 48h prior to use, endogenous substrates would be depleted,

expecially since steroid lack would inhibit protein breakdown and thus

limit the availability of amino acids. Hydrocortisone treatment would cause

breakdown of protein in muscle (Long et al., 1940), and the amino acids

released would stimulate insulin secretion, (Floyd et al., 1966),

204

as well as providing a carbon source for gluconeogenesis. The

facilitation of glycogenesis by the hexoses was therefore probably

due to the fact that they were present initially in the pre-treatment

in the presence of hydrocortisone, so that insulin secretion could be

stimulated immediately (and possibly potentiated by fructose). A

specific role for fructose in the restoration of synthesis was not however,

investigated.

(b) The role of insulin in hepatic glycogen accumulation in

adrenalectomised rats. Several series of experiments have suggested that

glucocorticoid - activated glycogenesis is mediated by insulin (see

above) and that the impairment in hepatic glycogen accumulation in

adrenalectomised rats is a consequence of insulin deficiency. The present

experiments confirm this suggestion, in that insulin (plus hexoses to

prevent fatal hypoglycaemia) administered to the intact animal, restored

. the capacity for net hepatic glycogen synthesis. The rate obtained was

identical to that found after hydrocortisone and hexose treatment.

At 'first sight, it would be paradoxical for insulin deficiency

to provide the sole explanation for the total loss of glycogen synthetic

capacity since the defect in starved diabetic animals is less severe.

However, it is possible that insulin lack in 48h-starved adrenalectomised

rats is indeed more severe than in the matched diabetic animals used

previously (they were only moderately diabetic and did not require insulin

maintainence). On starving diabetic animals, glycogen mobilisation is

slow (Friedmann, et al., 1963) and hyperglycaemia correspondingly slow

to disappear; thus insulin may still be detected in blood of starved

streptozotocin-diabetic rats (Junod et al., 1969; Schein et al., 1971 )

whereas the concentration in starved adrenalectomised rats is very low

(Van Lan et al., 1974).

205

A near total lack of circulating insulin would not matter in

the starved adrenalectomised state since without adrenal glands serious

ketonaemia would not supervene, adrenal gland hormones having a cofactor

role in the provision of substrate for hepatic ketogenesis (Chernick et al 1972).

However, since negligible rates of synthesis were obtained

after insulin in vivo for ih, it would seem unlikely that the metabolic

changes due to adrenalectomy are only a result of insulin deficiency.

It would also appear that an unknown factor (or factors) is implicated in

the restoration of glycogen accumulation (which could be a site of lesion

in adrenalectomy), since no direct hepatic effect of insulin was observed.

4.3.3 The characteristics of glycogen synthetase and

phosphorylase in the perfused liver of


The present experiments suggest that the impairment in hepatic

glycogen synthesis is due to an impairment in the responsiveness of both

hepatic glycogen synthetase and phosphorylase to substrates. Thus during

the perfusion of livers from adrenalectomised rats, in the absence of

pre-treatment in;vivo for more than lh, there was an impairment in the

increase in activity of glycogen synthetase "a" and fall in phosphorylase

seen in response to glucose and 03-substrates in the normal starved'

animal. A similar impairment in glycogen synthetase was observed in the

diabetic rat,although it was not as complete as the enzyme responded to

fructose (for Discussion see Section 4.2.). .

Loss of control of both glycogen synthetase and phosphorylase

by glucose has been previously reported in the perfused liver of 18h-

starved adrenalectomised rats (Miller et al., 1975). These enzymes did

however, respond normally in the fed adrenalectomised rat. A comparable

difference between the fed and starved state was observed in the activity

of glycogen synthetase phosphatase; the enzyme was reduced in the fasted

206 •

rat but not in the fed (Glinsmann, et al., 1970; Gruhner & Segal, 1970;

Mersmann & Segal, 1969). These results could implicate insulin in the

maintainence of glycogen synthetase and phosphorylase activity.

The action of steroids on hepatic glycogen phosphorylase activity

is not completely clear. As has been shown, the response of phosphorylase

to substrates (i.e., inactivation) is impaired on adrenalectomy (see also

Miller et al., 1973). This result suggests glucocorticoids act on

phosphorylase phosphatase, which is known to increase after steroid

administration in vivo (Stalmans et al., 1970). However, the activation

of hepatic phosphorylase by glucagon (Saitoh & Ui, 1975), or adrenalin

and cyclic AMP (Schaeffer et al., 1969),is lost on adrenalectomy,

implicating steroids in the conversion of "inactive" phosphorylase to

the "active" form i.e., at phosphorylase kinase. Measurements of hepatic

phosphorylase activity in the adrenalectomised rat have shown normal

levels of "active' enzyme, although there is a reduction in the "inactive"

form (Schaeffer et al., 1969). This reduction in activity of "inactive"

phosphorylase explains the lack of response of the enzyme to glucagon,

adrenalin and cyclic AMP. It would therefore appear that adrenal cortical

steroids are necessary for the maintenance of enzyme activity and are

involved in both the activation and inhibition of hepatic phosphorylase

activity.

Overall, glacocorticoids appear to be implicated in both sides of

glycogen turnover, in affecting the activities of both glycogen synthetase

and phosphorylase, in a manner which is not fully clear.

It has already been shown that the metabolic alterations in

hepatic glycogen metabolism due to adrenalectomy are not reversed rapidly,

even with hydrocortisone in vivo, and since there appears to be a total

impairment in the responsiveness of the enzymes glycogen synthetase and

phosphorylase, adrenal corticosteroid deficiency maybe considered as a

207

more severe state than diabetes.

Taken overall, the present results are compatible with the

notion that circulating insulin levels control the capacity for

hepatic glycogen synthesis, in relation to the dietary state, and to

pancreatic or adrenal endocrine status. This insulin effect appears

not be be di-rect on the liver, and its nature remains to be elucidated.

A similar problem exists for the short-term role of insulin in hepatic

ketogenesis (McGarry et al., 1.975).

208

4.4 THE ROLE OF THE HORMONES OF THE POSTERIOR-.

PITUITARY GLAND IN HEPATIC CARBOHYDRATE METABOLISM

4.4.1 Theos ard hormones and

the metabolism of liver carbohydrate.

The experiments described above (Section 3.3) show

that (8 -arginine) -vasopressin can stimulate the breakdown of liver

glycogen. This confirms the results obtained with higher concentrations

of vasopressin in liver slices (Heidenreich et al., 1963) and in the

perfused liver (Vaisler et al., 1965a) and the inference that was drawn

from experiments in vivo in which vasopressin was injected into the

hepatic portal vein (Bergen et al., 1960). Inhibition of glycogen

accumulation was observed in the perfusion and during intravenous

infusion'of vasopressin in vivo, showing that glycogen turnover (i.e.,

synthesis and breakdown) is affected by the hormone, and also that

vasopressin can affect hepatic glycogen metabolism in the intact animal.

It would seem likely that this prevention of glycogenesis in the starved

rat is due to the same action of vasopressin as in the atimulation of

glycogenolysis in the fed rat.

The effects of (8-arginine)-vasopressin observed in

the perfused rat liver were obtained with concentrations which can occur.

in vivo during "shock" or "stress" (see. Ginsburg, 1968), but especially

during acute haemorrhage (Forsling et al., 1971; Ginsburg & Heller, 1953).

Similar concentrations are necessary for the pressor action of the

hormone, exerted on extrahepatic tissues (see Altura, 1970; Dekanski,

1952). It is unlikely however, that the blood levels of vasopressin

in the unstressed animal are high enough to affect the liver. This may

also be the situation in regard to adrenalin action.

209

The action of vasopressin on hepatic carbohydrate metabolism

may be relevant to states of "shock" or "stress" for two different

reasons. Firstly, tissues require glucose during "stress", and the

present results show that vasopressin may contribute to the hyperglycaemia

usually associated with these states of "shock" (see Johnson, 1972),

by activating glycogenolysis and increasing gluconeogenesis. Secondly,

there are large quantities of water stored in conjunction with glycogen,

amounting to 2-4 times the weight of glycogen (Fenn, 1939). In a fed

rat of about 200g the hepatic glycogen content may be as much as ig and

the associated water up to 3m1. It is known that this water can be

mobilised e.g., during perfusion (Mortimore, 1961) or starvation (Herrera

& Freinkel, 1968). Therefore on stimulation of glycogenolysis by

vasopressin this hepatic water (which could amount to as much as one-third

of the plasma volume) would be released into the blood, without

prejudice to organ function. This action of vasopressin, in the

maintenance of plasma volume, would be especially relevant during, for

example, haemorrhage. During hypovolaemic stress, the hyperglycaemia

would serve to maintain extracellular fluid volume (including blood),

by osmotic action. These actions of vasopressin are therefore consistent

with the accepted role of the hormone in the preservation of plasma

volume, composition and pressure.

More generally, the action of vasopressin on the liver represents

a third main function of this hoimione in the rat, in addition to its

antidiuretic and pressor effects.

Since (8-arginine)-vasopressin caused glycogen breakdown, and

also moderate stimulation of gluconeogenesis (in livers from starved rats

that were perfused in the absence of added glucose), its actions on the

liver resemble those of glucagon (Exton et al., 1970). The minimum

210

effective circulating concentrations of these hormones (in the rat)

are of the same order, i.e., about 100pg/M1, as 11. units of (8-arginine)

-vasopressin corresponds to 2.5pg (for discussion of hepatic sensitivity

to glucagon see &ton et al., 1970; Sokal, 1966). As with glucagon (Pauk

& Reddy, 1971), the liver of the starved rat appeared to be more

sensitive to vasopressin than the liver of the fed rat.

The results obtained with oxytocin confirm those obtained

in vitro by other workers (Heidenreich et al., 1963; Vaisler et al.,

1965b), although Heidenreich et al., (1962) found no effect of oxytocin

in vivo, on the blood glucose concentration of the rat (presumably

reflecting the insensitivity of the liver to this hormone). Since

massive doses of the hormone were required to inhibit glycogen

accumulation in the perfusion, this action probably has no physiological

significance.

Hepatic and vascular responses to hormones of the posterior-

pituitary gland may be compared. The, two tissues exhibit (i) comparable

sensitivity to (8-arginine)-vasopressin (both responses .being much less

sensitive than that of the kidney) and (ii) insensitivity to oxytocin.

These observations suggest that the receptors to vasopressin in blood

vessels and liver may resemble each other in their characteristics, but

perhaps do not resemble the renal receptor to the hormone.

4.4.2 The mechanism of vasopressin action

In the kidney, vasopressin can stimulate glycogen breakdown

(Darnton, 1967) and gluconeogenesis (Stumpf et al., 1972). Since both

the renal action of vasopressin (Beck et al., 1971) and the hepatic

action of glucagon and possibly adrenalin (Exton et al., 1970) may be

211

mediated by an increase in the tissue concentration of cyclic AMP,

it appeared possible that the action of vasopressin on liver

carbohydrate metabolism would be similarly mediated. However, hepatic

cyclic JP does not rise in response to vasopressin (Kirk and Hems,

1974), so that a different "second messenger" may be implicated, in

vasopressin action.

A small increase in hepatic phosphorylase activity was

observed when vasopressin was administered in vivo. The activation of

the enzyme was not as significant as that obtained with glucagon, under

the same conditions, although the response and time course of activation

were similar to those obtained with adrenaline This provides a further

similarity in the actions of vasopressin andadrenalin on the rat liver

in addition to the lack of a role of cyclic AMP; thus the increase in

this "messenger" due to adrenalin, in comparison with that produced

by glucagon, is small (Kirk & Hems, 1974). It has been suggested that

adrenalin action on hepatic glucose output (cc receptor-mediated effect)

probably does not involve cyclic AMP as a messenger (Sherline et al.,

1972; Tolbert & Fain, 1974; Tolbert et al. 1975).

An observed increase in hepatic phosphorylase activity could

in general be due to either of two mechanisms : an increase in protein

kinase or a decrease in phosphorylase phosphatase activity. However,

in keeping with its lack of effect on hepatic cyolic AMP, vasopressin

(administered in vivo) does not activate hepatic protein kinase activity

(Keppens & DeNulf, 1975). Glucagon (Keppens & DeWulf, 1975) and

adrenalin (see Butcher et al., 1972) do, however, increase protein kinase,

the rise being more marked with glucagon than with adrenalin; this is in

agreement with the different increases in cyclic AMP concentrations

known to be produced by these hormones (Exton et al., 1971). The effect

212

of vasopressin on phosphorylase phosphatase has not been assessed.

It is known however/ that stimulation of the splanchnic nerve causes

activation of hepatic phosphorylase without any increase in cyclic AMP

(Shimazu & Amakawa, 1968 & 1975). It has been shown that the activity

of phosphorylase kinase is not altered, but that the activity of

phosphorylase phosphatase decreases promptly and markedly, on

splanchnic nerve stimulation. Glucagon and adrenalin have no significant

effect. From such circumstantial considerations,it seems plausible that

vasopressin could increase hepatic phosphorylase activity by inhibiting

phosphorylase phosphatase. This remains to be tested.

The metabolic actions of both vasopressin and adrenalin can

occur in the absence of any change in total hepatic blood flow, but it

may be that an explanation for the activation of phosphorylase by these

hormones lies in hypoxia produced by redistribution of blood flow, away

from the periphery of the lobes. Such a conclusion was drawn from

experiments showing that adrenalin inhibited hexobarbitone metabolism

by the perfused liver although it did not in rat liver slice studies

(Boobis & Fowls / 1974).

Glucagon, adrenalin and vasopressin are hormones which cause

hepatic glycogenolysis by increasing phosphorylase activity, and increase

gluconeogenesis. It does appear however, that although they have common

effects they all act via different mechanisms :. glucagon acting on

protein kinase activity by increasing cyclic AMP/ vasopressin possibly

decreasing phosphorylase phosphatase activity, and adrenalin acting via

local

hypoxia (cyclic AMP independent route). Many aspects of these

actions clearly still require elucidation.

213

REFERENCES

Akpan, J.0., Gardner, R.& Wagle, S.R. (1974) Biochem. Biophys. Res. Commun. 61, 222

Altura, B.M. (1970) Amer. J. Physio1.211, 222

Appleman, M.M., Krebs, E.G. & Fischer, E.H. (1966) Biochemistry 5, 2101

Ashmore, J., Stricker, F., Love, W.C. & Kilsheimer, G. (1961) Endocrinology 68, 599

Assal, J.P., Levrat, R., Stauffacher, W.& Renold, A.E.. (1971) Metab. Clin. Exp. 20, 850

Baker, N., Chaikoff, I.L. & Schusdek, A. (1952) J. Biol. Chem. 194, 435

Ballard, F.J. & Oliver, I.T. (1964) Biochem. J. 92, 131

Baron, D.N. & Roberts, P. (1963) J. Physiol. (London) 165, 219

Beck, N.P., Kaneto, T., Zor, U., Field, J.B. & Davis, B.B. (1971) J. Clin. Invest. 50, 2461

Bergen, S.S., Sullivan, R., Hilton, J.G., Willis, S.W. & Van Itallie, T.B. (1960) Amer. J. Physiol. 129, 136

Birch, G.G., Lee, E.Y.C. & Hems, D.A. (1974) Int. J. Biochem, 5, 867

Bishop, J.S. (1970) Biochem. Biophys. Acta 208, 208

Bishop, J.S. & Lamer, J. (1967) J. Biol. Chem. 242, 1354

Bishop, J.S. & Larner, J. (1969) Biochem. Biopys. Acta 171, 374

Bishop, J.S., Goldberg, N.S. & Lamer, J. (1971) Amer. J. Physiol. 220, 499

Blatt L.M. & Kim, K-H.(1971a) J. Biol. Chem. 246, 4895

Blatt, L.M. & Kim, K-H.(1971b) J. Biol. Chem. 2462. 7256

Bloxam, D.L. (1971) Br. J. Nutr. 26, 393

Boissonnas, R.A., Guttmnnn, St., Berde, B.& Konzett, H. (1961) 12ERILLI, 377

Boobis, A.R. & Fowls, G.(1974) Biochem. Pharmacol. 23, 3377

Burn, J.H. (1923) J. Physiol. London) 6/1, 324

Buschiazzo, H., Exton, J.H. & Park, C.R. (1970) Proc. Nat. Acad. Sci. U.S.A. 65, 383

214

Butcher, R.W., Robi.son, G.A. & Sutherland, E.W. (1972) in: Biochemical Action of Hormones (G. Litwack ed.) Vol. 2, p.219 Academic Press, New York and London.

Cahill, G.F., Hastings, A.B., Ashmore, J.& Zottu, S .(1958) J. Biol. Chem. 230, 125

Cahill, G.F., Ashmore, J., Renold, A.E. & Hastings, A.B. (1959) Amer. J. Med. 26, 264

Cash, W.D. & Kaplan, M.H. (1964) EndocrinolUY 74, 803

Cerasi, E. & Luft, R. (1970) Horm. Metab. Res. 2, 246

Chernick, S.S. & Chaikoff, I.L. , 1951) J. Biol. Chem. 188, 389 Chernick, S.S., Clark, C.M., Gar =ex, R.J. & Scow, R.U.C1972) Diabetes Clark, G.A. (1928) J. Physiol. (London) 64, 324 21, 946

Clark, D.G., Rognstad, R.& Katz, J. (1974) J. Biol.222Chen42, 2028

Claude, H.& Baudouin, A. (1912) C.R. Soc. Biol. 7,3, 732.

Coore, H.G. & Randle, P.J. (1964) Biochem. J1 93L 66

Cori, G.T. & Cori, C.F. (1940) J. Biol. Chem. 135, 733

Cori, C.F.,Cori, G.T. & Green, A.A. (1943) J. Biol. Chem. 151, 31

Curry, D.L., Curry, K.P. & Gomez, M. (1972) Endocrinology •12 1493

Darnton, S.J. (1967) 211aELI.jup. Physiol. 52, 392

Das, I.& Hems, D.A. (1974) Horm. Metab, Res. 6, 40

Dekanski, J. (1952) Brit. J. Pharmacol..1, 567

DeWulf, H. (1971) The Control of Glycogen Metabolism. Ph.D., Thesis. Vander, Louvain

DeWUlf, H. & Hers, H.G. (1967a) Ear. J. Biochem. 2, 50

DeWalf, H. & Hers, H.G. (1967b) Ear. J. Biochem. 2, 57

DeNUlf, H. & Hers, H.G.- (1968a) Ear. J. Biochem, 6, 552

DUalf, H. & Hers, H.G.: (1968b) Ear. J. Biochem. 6, 558

DeWalf, H., StaImans W.& Hers, H.G. (1968) Ear. J. Biochem. 6, 545

DeValf, H., Stalmans, W.& Hers, H.G.. (1970) Ear. J. Biochem. 15,, 1

DiPetro, D.L., Sharma, C.& Weinhouse, S. (1962) Biochemistry. 1, 455

Dunnigan, M.G. & Ford, J.A. (1975) Endocrinol. Metab. 40, 629

Eisenstein, A.B., Spencer, S., Flatness, S.& Brodsky, A. (1966) Endocrinology 79,.182

215

Exton, J.H. & Park, C.R. (1965) J. Biol. Chan. 240, 955

Exton, J.H. & Park, C.R. (1957) J. Biol. Chem. 242, 2622

EXton, J.H. & Park, C.R. (1968) J. Biol. Chem. 243, 4189

Exton, J.H., Mallette, L.E., Jefferson, L.S., Wong, E. H-A., Friedmann, N., Miller, T.B. & Park, C.R. (1970) 2.,...ecentPiorm. Res. 26, 411

Exton, J.H., Robison, G.A., Sutherland, E.W. & Park, C.R. (1971) J. Biol. Chem. 2461 6166

Exton, J.H.9 Friedmann, N., Wong, E.H-A., Brineaux, J.P., Corbin, J.D.. &.Park, C.R. (1972a) J. Biol. Chem. 24, 3579

Exton, J.H., Corbin, J.G. & Harper, S.C. (1972b) J. Biol. Chem. 247, 4996

Exton, J.H., Harper, S.C., Tucker, A.D. & Ho, R-J. (1973a) Biochem. Biophys. Acta. 329, 23

Exton, J.H., Harper, S.C., Tucker, A.L., Flagg, J.L. & Park, C.R. (1973b) Biochem. Biophys. Acta 329, 41

Fenn, W.O. (1939) J. Biol. Chem. 128, 297

Fischer, E.H., Pocker, A. & Saari, J.C. (1970) Essays in Biochemistry 6, 23

Floyd, J.C., Fajans,S.S., Conn, Knopf, R.F. & Hull, J. (1966) J. Clin. Invest. 45, 1487

Forsling, M.L., Martin, M.J. & Burton, A.M. (1971) J. Endocrinol, 51, 413

Friedmann, B., 238, 2899

Friedmann, B., 3729

Goodman, E.H. & Weinhouse, S. (1963) J. Biol. Chem.

Goodman, E.H. & Weinhouse, S. (1965) J. Biol. Chem. 210

Friedmann, B., Goodman, E.H. & Weinhouse, S. (1967) Endocrinology 81, 486

Friedmann, B., Goodman, E.H. & Weinhouse, S. (1970) Endocrinology 86, 1264

Gilboe, D.P. & Nuttall, F.Q., (1973) Biochem. Biophys. Res. Commun. a, 164

Gilboe, D.P. & Nuttall, F.Q. (1974) Biochem. Biophys. Acta 338, 57

Ginsburg, M. (1968) Handb. EXp. Pharmacol. 23, 286

Ginsburg, M.& Heller, M. (1953) J. Endocrinol. 9, 274

Glinsmann, W.H. & Hern,E.P. (1969) Biochem. Biophys. Acta 36, 931

Glinsmann, W.H. & Mortimore, G.E. (1968) Amer. J. Physiol. 215, 553

216

Glinsmann, W.H., Hern, E.P. & Lynch, A. (1969) Amer. J. Physiol. 216, 698

Glinsmann, W.H., Pauk, G. & Hem, E./1(1970) Biochem. Biophys. Res. Commun. 39, 774

Gold, A.H. (1970a) Biochemistry 9, 946

Gold, A.H. (1970b) J. Biol. Chem. 245, 903

Gold, A.H. & Segal, H.L. (1966) Mol. Parmacol. 2, 84

Good, C.A., Kramer, H. & Somogyi, M. (1933) J. Biol. Chem. 100, 485

Grodsky, G.M., Batts, A.A., Bennett, L.L., Vbella, C., McWilliams, N.B. & Smith, D.P. (1963) Amer. J. Physiol. 205, 638

Gruhner, K. & Segal, H.L. (1970)1.BI ta222Biochen , 508

Haft, D.E. (1967) Amer. J. Physiol. 213, 219

Haft, D.E. (1968) Diabetes 17, 244

Heidenreich, 0., Kook, Y.& Reus, E. (1962) Naunyn-Schmiedeberg's Arch. Exp. Pathol. Pharmakol. 243, 136

Heidenreich, 0., Kook, Y., Baumeister, L. & Keller, P. (1963) Naunyn-Schmiedebergls Arch. Exp. Pathol. Pharmakol. 245, 321

Hems, D.A. & Brosnan, J.T. (1970) Biochem. J. 120, 105

Hems, R., Ross, B.D., Berry, M.N. & Krebs, H.A. (1966) Biochem. J. 101, 284

Hems, R., Stubbs, M. & Krebs, H.A. (1968) Biochem. J. 107, 807

Herrera, E. & Preinkel, N. (1968) Biochem. Biophys. Acta 170, 244

Hers, H.G., Deliulf, H., Stalmans, W & Van den Berghe, G. (1970) Advan. Enzyme Regu1. 8, 171

Hildalgo, J.L. & Rosell-Perez, M. (1971) Rev. Esp. Fisiol. 27, 343

Hizukuri, E. & Larner, J. (1964) Biochemistry 3, 1783

Hohorst, H.J. (1963) in: Methodsof H.U., ed.) p266, Academia Press, New York

Hornbrook, K.R. (1970) Diabetes 19, 916

Hornbrook, K.R., Burch, H.B. & Lowry, 0.H. (1965) Biochem. Biophys. Res. Commun. 18, 206

Hornbrook, Burch, H.B. & Lowry, 0.H. (1966) Nol. Pharmacol. 22. 106

217

Hostmark, A.T. (1973) Acta physiol. Scan& 88, 248

Illiano G., Tell, G.P.E., Siegel, M.I. & Cuatrecasas, P. (1973) Proc. Nat. Acad. Sci. U.S.A. 70, 2443

Imrie, C.G. (1929) J. Ph. siol. (London) 67, 264

Jeffcoate, S.L. & Moody, A.J. (1969) Diabetologia. 5, 293

Jefferson, L.S., Exton, J.H., Butcher, R.W., Sutherland E.W. & Parks C.R. (1968) J. Biol. Chem. 243, 1031

Jeffrey, P.L., Brown, D.H. & Illingworth Brown, B. (1970) Biochemistry 9, 1403

John, D.W. & Miller, L.L. (1969) J. Biol. Chem. 244 6134

Johnson, I.D.A. (1972) Advan. Clin.2221:15, 255 Junod, A., Lambert, A.E., Stauffacher, W. & Renold, A.B. (1969) J.

Clin. Invest. 48, 2129

Keppens, S. & DeWulf, H. (1975) F.E.B.S. Letters_21, 29

Kirk, C.J. & Hems, D.A. (1974) F.E.B.S. Letters .AZ, 128 Kirk, C.J., Verrinder, T.& Hems, D.A. (1975) Biochem. Soc. Trans.

.3.2. 515

Krebs, H.A. & Henseleit, K. (1932) itopzzSELefsaphmi.o1z....Chem. 11-22' 33

Krebs, H.A.t Dierks, C. & Gascoyne, T. (1964) Biocham.11....21 112 Kreutner, W. & Goldberg, N.D.(1967) Proc. Nat. Acad. Sci. U.S.A. ..1515

Landau, B.R. (1965) Vitam. Horm. New York) 23, 1

Landgraf, R., Kotler-Brajtburg, J.& Hatschinsky, F.M. (1971) Proc. Nat. Acad. Sci. U.S.A. 8, 536

Larner, J. & Villar-Palasi, C. (1971) Current Topics in Cell. Reg. 3, 195

Lee, E.Y.C. & Whelan, W.J. (1966) Arch. BiochenL..131ophya...116, 162

Lejeune, N., Thines-Sempoux, D.& Hers, H.G. (1963) Biochem. J. 86, 16

Leloir, L.F., (1964) in: Ciba Foundation Suiposium on Control of Glycogen Metabolism (W.J. Whelan & M.P. Camerc7r7e77777.18 Churchill, London

Leloir, L.F. (1967) E-LLsp3229LD....isii.Monorah2E2 1, 3

Leloir, L.F. & Carlini, C.F. (1957) J. Amer. Chem. Soc. 79, 6340

Leloir, L.F. & Goldemberg, S.H. (1960) J. Biol. Chem, 23, 919

218

Little, J.B., Klevay, L.M.,Radford, E.P. & McGandy, R.P. (1966). Amer. J. Physiol. 211, 786

Levine, R. & Fritz, I.B. (1956) Diabetes 5, 209

Long, C.N.H., Katzin, B. & Fry, E.G. (1940) Endocrinology 26, 309

Longley,R.W., Bortnick, R.I. & Roe, J.H. (1957) Proc. Soc. Exp. Biol. Med. 94, 108

MacDonald, M.J.,Ball, T.N.Lauris, V. & Steinke, J. (1975) Biochem. 188

Mackrell, D.J. & Sokal, J.E. (1969) Diabetes 18, 724

Maddaiah, V.T. & Madsen, N.B. (1966) J. Biol. Chem. 241, 3873

Maddaiah, V.T. & Madsen N.B. (1968) Can. J. Biochem. 46, 521

Madison, L.L. (1969) Archt ZE:berIrlad.123" 284

Mahler, R.J. & Szabo O. (1970) latLLIJILD1L Eat 11, 271

Malaisse, W.J., Malaisse-Lagae, F.,McCraw, E.F. & Wright, P.H. (1967) Proc. Soc. E2g. Biol. Med. 124, 924

Nanette, L.E., Exton, J.H. & Park, C.R. (1969) J. Biol. Chem. 244, 5724

Mansford, K.R.L. (1967) Diabetes 16, 475

Marsh, W.H., Fingerhut, B. & (1965) 21122 21.,..ien21.222, 624

McGarry, J.D. Wright, P.M. & Foster, D.W. (1975) J. Clin. Invest. Z,. 1202

Merlevede, W., Goris, J. & DeBrandt, C. (1969) Ear. J. Biochem. ILL 499

Mersmann, H.J. & Segal, H.L. (1967) Proc. Nat. Acad. Sci. U.S.A. 211. 1688

Mexsmann, H.J. & Segal, H.L. (1969) 1701

Miller, L.L.,Bly, C.G., Watson, M.L. & Bale, W.F. (1951) L...}22,c laa,21, 431

Miller, M., Drucker, W.R., Owens, J.E., Craig, J.W. & Woodward, H. (1952) J. cliilLaampIl 51, 115

Miller, T.B. & Larner, J. (1973) J. Biol. Chem. 248, 3483

Miller, T.B., Hazen, R. & Larner, J. (1973) Bi_o12111.Bi.ophystRes. Common. 7, 466

Mills, G.T. & Smith, E.E.B. (1963) in: Methods of Ersti-acAnalys1- (Bergmeyer, H.U. ed. ) P . 581, Academic Press, London

Mondon, C.E. & Burton S.D. (1971) Amer. J. phapiol. 220, 724

219

Mondon, C.E. & Mortimore G.E. (1967) 173

Moody, A.J., Jeffcoate, S.L. & Volund, A. (1970) Horm. Metab. Res. 22. 193

Mortimore, G.E. (1961) Amer. J. Physiol. 201 1315

Nichols, W.K. & Goldberg, N.D. (1972) BiocheiBiohm"42-22ba2, 245

Nordlie, R.C. (1968) in: Control of Glycogen Metabolism (W.J. Whelan ed.) Universitets for laget, Oslo, p. 1531Academic Press, London,

Nuttall, P.Q., (1972) in: Handbook of Physiolam, Section 7 (Endocrinology, Greep, R.O. & Astwood, E.B., eds.) Vol. 1. p.395, American Physiological Society.

Olavarria,J.M., Godeken, 0.G.R., Sandruss, R. & Flawia, M. (1968) Biochem. Biophys. A2ta165, 185

Porter, K.R. & Bruni, C. (1959) CancerEu: la, 997

Pauk, G.L. & Reddy, W.J. (1971) Diabetes 20, 129

Pilkis, S.J. & Park, C.R. (1974) Ann. Rev. Pharmacol.....1i, 365

Pilkis, S.J., Exton, J.H., Johnson, R.A. & Park, C.R. (1974) Biochem. Biophys. Acta 343, 250

Rall. T.W. & Sutherland, E.W. (1958) J. Biol. Chem. 212., 1065

Randle, P.J. & Hales, C.N. (1972) in: HandbaokaLEILliala, Section 7 (Endocrinology, Greep, R.O. & Astwood, E.B. eds.) Vol. 1 p.219,American Physiological Society

Renold, A.E., Teng, C-T., Nesbett, F.B. & Hastings, A.B. (1953) J. Biol. Chem. 204, 533

Renold, A.E., Hastings, A.B., Nesbett, F.B. & Ashmore, J. (1955) J. Biol. Chem. 213, 135

Renold, A.E., Ashmore, J. & Hastings, A.B. (1956) Vitam. Horm. New York) 14, 139

Ross, B.D., Hems. R., Freedland, R.A. & Krebs, H.A. (1967) Biochem. 869

Ruderman, N.B. & Herrera, M.G. (1968) .limer:L.layIsio1„...2141 1346

Ryman, B.E. & Whelan, W.J. (1971) Advan. Enzvmol. Rel. Areas Mol. Biol. 34, 285

Saitoh, Y. & Ui, M. (1975) B5ochem. Biophys. Acta 404.2. 7

Salas, M., Vinuela, E. & Sols, A. (1963) .L1uLshm,2322 3535

Salmon, D.M.W., Bowen, N.L. & Hems, D.A. (1974) Biochem. J. 142, 611

Salsas, E. & Lamer, J. (1975) J. Biol. Chem. 250, 1833

220

Schaeffer, L.D., Chenoweth, M. & Dunn, A. (1969) Biochem. Biophys. Acta 192, 292

Scharff, R. & Wool, I.G. (1966) Biochem. J. !II, 173

Schein, P.S., Alberti, K.G.M.M. & Williamson, D.H. (1971) Endocrinology 8g, 827

Schillinger, E., Loge, E., Schroder, E., Klieger, E. & Lubke, K. (1972) EuIt Bj.ocl11.2 473

Schimassek, H. (1963) 131.ochenit21 336, 460

Schimassek, H.& Gerok, W. (1965) Biochem. Z. 343, 407

Seglen, P.O. (1974) Biochem. Biomes. Acta M, 317

Sharma, C., Manjeshwar, R. & Weinhouse, S. (1963) J. Biol. Chan. 238, 3840

Sherline, P., Lynch, A. & Glinsmann, W. (1972) Endocrinology21, 680

Shimazu, T. & Amakawa, A. (1968) Biochem. Blophys. Acta 165, 335

Shimazu, T & Amakawa, A. (1975) Biacilein.33i2=stAs-La325...2„ 242

Soderling, T.R. & Park, C.R. (1974) Advan.aclic Nucleotide Res. AL 284

Sokal, J.E. (1966) Endocrinoloy 78, 538

Sokal, J.E., Miller, L.L. & Sarcione, E.J. (1958) Amer. J. Physiol.115.9 295

Sokal, J.E., Sarcione, E.J. & Henderson, A.M. (1964) Endocrinology 930

Soskin, S. (1941) Physiol. Rev. 214 140

Soskin, S., Essex, H.E. Herrick, J.P. & Mann, F.C. (1938) Amer. 11:21-asj,..2LL2.1.2.. 558

Stalmans W., BeWulf, H., Lederer, B. & Hers, E.G. (1970) Eur. J. Biochem. 151. 9

Stalmans, DeWulf,H.& Hers, H.G. (1971) 582 Stalmans, W., DeWuif, H.,Hue, L..& HersiII.G.(1974a) Eur. J. Biochem. 41, 127

Stalmans, W., Laloux, M. & Hers, H.G. (1974b) Eur. J. Biochem. Ai. 415

Steele, R. (1966) Eraeb. PhzE121. Biol. Chem. Exp. Pharmakol. 57, 91

Stehle, R.L. (1950) ItiVitam.Hor c8, 216

Steiner, D.F. (1964) Nature (London 204, 1171

221

Steiner, D.P. (1966) Vitam. Horm._.(New ysistl21.2. 1

Steiner, D.F. & King, J. (1964) J. Biol. Chem. 239, 1292

Steiner, D.F., Rauda, V. & Williams R.H. (1961) J. Biol. Chem. 216z.299

Stetten, De.W & Boxer, G.E. (1944) 21131-01tSL511.169 271

Stumpf, B., Bole, A. & Seubert, W. (1972)n.33529zBiochei Commun. 49, 164

Sutherland, E.W. & Wosilait, W.D. (1956) J. Biol. Chem. 218 459

Tan, A.W.H., & Nuttall, F.Q., (1974) Fed. Proc. .1311 1894A,

Tata, J.R., (1964) 2L2.0.11229 284

Thomas, A.J. (1969) in: Automation Mechanisation and Data Handling in Microbio Ed. by Soc. Applied Bacteriology, Technical Handbook. Academic Press.

Thurston, J.H.,Jones, E.M. & Hauhart, R.E. (1974) Diabetes 23 597

Tolbert, M.E.M. & Fain, J.N. (1974) J. Biol. Chem. 249 1162

Tolbert, M.E.M.,Butcher, P.R. & Fain, J.N. (1973) J. Biol. Chem. 248a 5686

Unger, R.H. (1972) Advan. Metab. Dis. 6, 73

Vaisler, L., Costiner, E. & Gheorghiu, C. (1965 ) St. Cercet. Endocrinol. 16, 403

Vaisler, L., Costiner, E. & Gheorghiu, C. (1965b) St. Cercet. Endocrinol. 16, 557

Van den Berghe, G., Hue, L. & Hers, H.G. (1973) Biochem. J. 214., 637

Van Lan, V., Yamaguchi, N., Garcia, M.J., Ramey, E.R. & Penhos, J.C. (1974) Endocrinology 2A, 671

Villar-Palasi, C. & Larner, J. (1960) Bi2...Llen....11 21 ta, 171

Vinuela, E., Salas, M. & Sols, A. (1963) lz Bio.238" 1175

Walaas, 0., Walaas, E. & Gronnerod, 0. (1974) -1,-9:UZda1212. 93

Walker, D.G. (1963) 13iochenys. Actl_71, 209

Walker, D.G. & Rao, S.(1964) Biochem. J. 90, 360

warn, A.K., Siebler, G., Zepf, E. & Schimassek, H. (1974) 52P.... Sela211g.....1Pilsen1- 7 59 353

222

Winternitz, LW., Long, C.N. & Dintzis, R. (1957) Endocrinology dla 724

Whelan, W.J. (1971) Biochem. J. 122, 609

Wolf, D.P., Fischer, E.H. & Krebs, E.G. (1970) Biochemistry 9, 1923

Wosilait, W.D. & Sutherland,E.W. (1956) J. Biol. Chem. 218, 469

Zaragoza-Hemmans, N. & Felber, J.P. (1972) Horm. Metab. Res. 4, 25

Documents

TED, ROLE OF HORMONES IN HEPATIC CARBOHYDRATE METABOLISM · 4.2.2 Hepatic glycogen accumulation in the starved diabetic rat 193 4.2.3 Properties of glycogen synthetase and phosphorylase