Somatostatin Receptor Type 2 (SSTR2) Antagonism and Hypoglycemia

in Diabetes

by

Jessica Tin Yan Yue

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Graduate Department of Physiology University of Toronto

© Copyright by Jessica Tin Yan Yue, 2011

ii

Somatostatin Receptor Type 2 (SSTR2) Antagonism and Hypoglycemia in Diabetes

Jessica Tin Yan Yue

Doctor of Philosophy

Graduate Department of Physiology University of Toronto

2011

Abstract

Hypoglycemia is one of the most serious acute complications in intensively treated diabetes.

Recurrent hypoglycemia predisposes individuals to subsequent hypoglycemia, and

diminished counterregulatory hormone responses increase this threat. Elevated pancreatic

and/or circulating somatostatin has been reported in diabetic humans and animals, and we

postulated that excessive somatostatin contributes to the attenuation of counterregulatory

hormone release during hypoglycemia in diabetes. It is known that somatostatin

suppresses stimulated secretion of glucagon, epinephrine, and corticosterone. We

hypothesized that selective somatostatin receptor type 2 (SSTR2) antagonism would:

(Study 1) improve hormone counterregulation to hypoglycemia, and (Study 2) ameliorate

hypoglycemia in recurrently hypoglycemic rats. Using both high (10 U/kg) and low (5 U/kg)

dose insulin to induce hypoglycemia, we demonstrate that inhibiting the action of

somatostatin on SSTR2 normalizes the severely attenuated glucagon and corticosterone

responses to acute hypoglycemia in diabetic rats. These improvements were specific to

iii

diabetes since SSTR2 antagonism did not increase these hormones in non-diabetic rats in

response to hypoglycemia. In the absence of hypoglycemia, SSTR2 antagonist neither

markedly alters glycemia nor causes sustained elevations in counterregulatory hormones in

diabetic animals. Diabetic rats exhibit up to 65% and 75% more pancreatic and plasma

somatostatin than non-diabetic rats following hypoglycemia, respectively. Despite

improvements of glucagon and corticosterone, expression of gluconeogenic enzymes

PEPCK1 and G6Pase was unaltered. SSTR2 antagonism reduced the glucose requirement

during a hypoglycemic clamp induced with a lower dose of insulin. In recurrently

hypoglycemic diabetic rats, we demonstrate that SSTR2 antagonist treatment reduces the

depth and duration of hypoglycemia and promotes the recovery to euglycemia, without

affecting the glycemia-lowering effect of insulin. This amelioration of hypoglycemia by

SSTR2 antagonism may be attributable in part to the observed modest improvements of

glucagon, epinephrine, and corticosterone counterregulation following recurrent

hypoglycemia. These results implicate an important role for increased pancreatic, and

possibly circulating, somatostatin in defective hypoglycemic counterregulation in diabetes.

iv

Acknowledgments

I would fist like to thank my supervisor, Dr. Mladen Vranic, and co-supervisor, Dr. Adria

Giacca, for their support and direction throughout my PhD studies. Dr. Vranic – it has truly

been an honour to have been mentored by you. You provided me with so many

opportunities to further my research, and I have learned so much as a result of your

teaching. You allowed me to be independent while providing me with your wise counsel. It

is a great privilege to have been your student. I will look back with much fondness on the

nine years spent in your laboratory, first as a third-year undergraduate student and

subsequently earning my Master’s and Doctoral degrees under your supervision. Adria –

your expertise, sincerity, and kindness have meant so much to me over the years. Your

genuine care and selfless giving of your time and knowledge have made a deep and lasting

imprint on me. I am extremely thankful to have had the both of you as my supervisors. I

extend my appreciation to the members of my supervisory committee, Dr. Daniel Drucker

and Dr. Qinghua Wang. Thank you for your guidance, your generous expertise, and your

encouragement. I am deeply grateful for your scientific insights and for all your help with

progressing my studies. You have demonstrated much kindness and have been very

generous with your time and advice.

I would like to acknowledge and thank Dr. Suad Efendic (Karolinska Insitute) and Dr. David

Coy (Tulane University) for their collaboration. I would like to thank Dr. Ian Stewart (The

Innovations Group, University of Toronto) for his help and expertise in filing our patent. I

would like to thank Judy Blumstock (MaRS Innovations) and Ian for introducing me to the

area of commercialization in research and for their help in connecting us with people

interested in seeing our hard work translated.

v

I would like to thank Elena Burdett and Loretta Lam not only for their excellent and

valuable technical assistance and knowledge, but also for their much appreciated support,

thoughtfulness and constant camaraderie over the years. Elena – I will always cherish your

friendship and the laughter we shared together. Thank you for taking such great care of me

and for involving me in your life. I would like to thank Dr. Holly Bates, Dr. Christine

Longuet, and members of the Drucker laboratory for their helpful advice and for sharing

their time and methods of RNA isolation RT-PCR with me. Holly – I truly treasure your

friendship and support and will take with me many happy memories of our time together.

To former project students who participated in this project – Eric Chan, Andrea Kokorovic,

Jamila Husein, Adam Sirek, Angela Chung, and Garry Thomas – thank you for all your help

and youthful inspiration! I am indebted to their hard work and dedication to make this

project succeed.

To Bryan – thank you for your constant optimism and your support of me. Thank you for

the late-night pick-ups at the lab, for bringing food, and for lending an extra set of hands on

the weekends when I needed them. Just for understanding – thank you. You enrich my life

in so many ways, and I am so appreciative of all your encouragement, patience, and love.

Lastly and most importantly, I would like to acknowledge my family for their unceasing

support and encouragement in all that I do, whether I deserve it not. To my sister and

brother-in-law, Josephine and Karwin – thank you for always helping to keep my spirits up

and for keeping me real. And especially to my parents, Suzanna and Edwin – I thank you

for your constant affirmation, your prayers, and your love. Thank you for continually

teaching me and reminding me to always keep my focus on the most important things in

life. Truly, the impact of the lessons I have learned at home far surpass the knowledge

acquired from my studies.

Jessica T.Y. Yue

vi

Table of Contents

 

Acknowledgments .................................................................................................. iv 

Table of Contents ................................................................................................... vi 

List of Tables .......................................................................................................... xi 

List of Figures ....................................................................................................... xii 

List of Abbreviations ............................................................................................. xv 

Chapter 1 General Introduction ...............................................................................1 

1.1  Hypoglycemia ................................................................................................ 1 

1.1.1  Normal sensing of hypoglycemia: sites of glucose detection ........................ 2 

1.1.2  Normal counterregulatory hormone responses to hypoglycemia .................. 4 

1.1.2.1  Insulin. ................................................................................... 4 

1.1.2.2  Glucagon. ................................................................................ 4 

1.1.2.3  Epinephrine. ............................................................................ 7 

1.1.2.4  Norepinephrine. ....................................................................... 8 

1.1.2.5  Growth hormone and glucocorticoids. .......................................... 9 

1.1.2.6  Glucose autoregulation. ........................................................... 10 

1.2  Hypoglycemia in diabetes .............................................................................. 11 

1.2.1  Clinical Significance – Hypoglycemia in type 1 diabetes mellitus ................ 11 

1.2.2  Clinical Significance – Hypoglycemia in type 2 diabetes mellitus ................ 12 

1.3  Defective counterregulation in diabetes ........................................................... 14 

1.4  Mechanisms of defective counterregulation in diabetes ...................................... 14 

1.4.1  Glucagon counterregulation in impaired in diabetes ................................. 15 

1.4.2  Epinephrine counterregulation in diabetes .............................................. 18 

1.4.3  Norepinephrine, growth hormone, and glucocorticoid counterregulation in diabetes ............................................................................................ 20 

1.4.4  Hypoglycemia unawareness in diabetes ................................................. 21 

1.4.5  Hypoglycemia-associated autonomic failure (HAAF) in diabetes ................. 22 

1.5  Recurrent hypoglycemia in non-diabetic subjects .............................................. 23 

1.6  Recurrent hypoglycemia in diabetes ................................................................ 23 

1.7  Somatostatin ............................................................................................... 25 

1.7.1  Functions .......................................................................................... 26 

1.7.1.1  General somatostatin effects in pancreatic islets. ........................ 26 

vii

1.7.1.2  Arguments against local somatostatin effects on pancreatic islets. . 27 

1.7.1.3  General somatostatin effects in the brain and pituitary gland. ....... 27 

1.7.1.4  General somatostatin effects in the gastrointestinal tract. ............. 28 

1.7.1.5  Other effects of somatostatin relating to glucose homeostasis. ...... 28 

1.7.2  Localization of somatostatin-producing cells ........................................... 29 

1.7.2.1  Pancreatic somatostatin-producing cells. .................................... 29 

1.7.2.2  Brain somatostatin-producing cells. ........................................... 30 

1.7.2.3  Gastrointestinal somatostatin-producing cells. ............................ 31 

1.7.2.4  Other somatostatin-producing cells. .......................................... 31 

1.7.3  Synthesis and structure ....................................................................... 31 

1.7.3.1  Cortistatin. ............................................................................ 32 

1.7.3.2  Neuronostatin. ....................................................................... 33 

1.7.3.3  Apelin ................................................................................... 33 

1.7.4  Regulation of somatostatin synthesis ..................................................... 34 

1.7.5  Regulation of somatostatin secretion ..................................................... 36 

1.7.5.1  Regulation of pancreatic somatostatin. ...................................... 36 

1.7.5.2  Regulation of brain somatostatin. .............................................. 37 

1.7.5.3  Regulation of gastrointestinal somatostatin. ............................... 37 

1.8  Somatostatin receptors (SSTRs) ..................................................................... 38 

1.8.1  General localization of SSTRs ............................................................... 39 

1.8.2  General functions of SSTRs .................................................................. 41 

1.8.3  Specific localization, ligand binding, and signalling of SSTR2 ..................... 42 

1.9  Somatostatin in type 1 diabetic humans and animal models ............................... 44 

1.9.1  Brain somatostatin in experimental diabetes .......................................... 47 

1.10  Somatostatin in type 2 diabetic humans and animal models.............................. 48 

1.11  Diabetes-induced changes in SSTR expression ................................................ 48 

1.12  Somatostatin: hypoglycemia-induced changes ................................................ 49 

1.12.1 Hypoglycemia-induced somatostatin and SSTR changes in the brain .......... 51 

1.13  Somatostatin and its analogues in diabetic therapy ......................................... 51 

1.14  Somatostatin antagonist – PRL-2903 ............................................................. 52 

Chapter 2 Objectives ............................................................................................. 54 

2.1  Specific Aim: Study 1 ................................................................................... 54 

2.2  Specific Aim: Study 2 ................................................................................... 55 

Chapter 3 General Materials and Methods ............................................................. 56 

3.1  Experimental animals ................................................................................... 56 

3.1.1  Care and maintenance ........................................................................ 56 

viii

3.1.2  Streptozotocin-diabetic rat model of diabetes mellitus ............................. 56 

3.1.3  Induction of diabetes using streptozotocin .............................................. 58 

3.1.4  Chronic vessel catheterization .............................................................. 59 

3.1.4.1  Pre-operative preparation. ....................................................... 59 

3.1.4.2  Surgical procedure. ................................................................. 59 

3.1.4.3  Post-operative care. ................................................................ 61 

3.1.4.4  Catheter patency and blood sampling. ....................................... 62 

3.2  General laboratory methods ........................................................................... 62 

3.2.1  Blood Glucose Determination ............................................................... 62 

3.2.2  Preparation of blood sample collection tubes .......................................... 63 

3.2.3  Radioimmunoassays (RIA) for plasma hormone levels ............................. 64 

3.2.3.1  Plasma corticosterone assay. .................................................... 65 

3.2.3.2  Plasma glucagon assay. ........................................................... 66 

3.2.3.3  Plasma catecholamine assay. ................................................... 68 

3.2.3.4  Plasma somatostatin assay. ..................................................... 70 

3.2.3.5  Plasma insulin assay ............................................................... 72 

3.2.3.6  Plasma growth hormone assay ................................................. 74 

3.2.4  Pancreatic protein determination .......................................................... 76 

3.2.4.1  Pancreas homogenization for protein isolation ............................. 76 

3.2.4.2  Sep-Pak protein extraction ....................................................... 76 

3.2.5  Liver mRNA expression quantification .................................................... 77 

3.2.5.1  Total RNA preparation. ............................................................ 77 

3.2.5.2  cDNA synthesis. ..................................................................... 78 

3.2.5.3  Polymerase chain reaction (PCR) of -actin. ................................ 79 

3.2.5.4  Real-time quantitative PCR. ..................................................... 80 

3.3  Preparation of SSTR2 antagonist and saline infusates ........................................ 82 

3.4  Statistical analysis ........................................................................................ 82 

Chapter 4 Study 1: Effect of SSTR2 antagonism on counterregulation to hypoglycemia .................................................................................................... 84 

4.1  Aims .. ....................................................................................................... 84 

4.2  Experimental methods .................................................................................. 85 

4.2.1  Study design ..................................................................................... 85 

4.2.2  Hypoglycemia experiments .................................................................. 86 

4.2.3  Control experiments: SSTR2 antagonist in the absence of hypoglycemia .... 87 

4.2.4  Effect of SSTR2 antagonist in non-diabetic rats during hypoglycemia ......... 87 

4.3  Results ....................................................................................................... 88 

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4.3.1  Daily blood glucose, body weight, and food intake ................................... 88 

4.3.2  Hypoglycemia clamp experiments ......................................................... 89 

4.3.2.1  Pilot study: using 10 U/kg insulin and 1500 nmol/kg/h SSTR2 antagonist. ............................................................................ 89 

4.3.2.2  Using 10 U/kg insulin and 3000 nmol/kg/h SSTR2 antagonist. ...... 89 

4.3.2.3  Using 5 U/kg insulin and 3000 nmol/kg/h SSTR2 antagonist. ........ 91 

4.3.3  Effect of SSTR2 antagonist in the absence of hypoglycemia ...................... 92 

4.3.4  Pancreatic glucagon and pancreatic and plasma somatostatin measurements ................................................................................... 93 

4.3.5  Liver PEPCK1 and G6Pase mRNA expression ........................................... 93 

4.3.6  Effect of SSTR2 antagonist in non-diabetic rats during hypoglycemia ......... 94 

4.4  Discussion ................................................................................................... 94 

4.4.1  Role for SSTR2 in the enhancement of glucagon counterregulation ............ 94 

4.4.2  Potential role for SSTR2 in insulin secretion? .......................................... 98 

4.4.3  Role for SSTR2 in the enhancement of corticosterone counterregulation ..... 99 

4.4.4  Epinephrine responses to hypoglycemia were unaffected ......................... 103 

4.4.5  SSTR2 antagonist demonstrated no effect on growth hormone ................. 104 

4.4.6  SSTR2 antagonism does not cause hyperglycemia or undesired stress ...... 106 

4.4.7  Glucose requirement during hypoglycemia ............................................ 108 

4.4.8  Expression of gluconeogenic enzymes .................................................. 109 

4.4.9  Effect of SSTR2 antagonism during hypoglycemia in non-diabetic rats ....... 112 

4.5  Study 1 Conclusions .................................................................................... 114 

Chapter 5 Study 2: Amelioration of hypoglycemia using SSTR2 antagonism ....... 136 

5.1  Aims .. ...................................................................................................... 136 

5.2  Experimental methods ................................................................................. 137 

5.2.1  Study design .................................................................................... 137 

5.2.2  Recurrent hypoglycemia treatment ...................................................... 137 

5.2.3  Partial overnight fasting prior to hypoglycemia experiments .................... 138 

5.2.4  Hypoglycemia on Experimental Days 1 and 2 ......................................... 138 

5.3  Results ...................................................................................................... 140 

5.3.1  Pilot Studies: How the present method was established .......................... 140 

5.3.2  Daily blood glucose, body weight, and food intake .................................. 141 

5.3.3  Glycemia during recurrent hypoglycemia treatment ................................ 142 

5.3.4  Basal blood glucose and plasma hormone levels after recurrent hypoglycemia treatment ..................................................................... 142 

5.3.5  Blood glucose and plasma insulin levels during Experimental Days 1 and 2 142 

5.3.6  Counterregulatory hormones levels during Experimental Days 1 and 2 ...... 143 

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5.4  Discussion .................................................................................................. 146 

5.4.1  Amelioration of hypoglycemia with SSTR2 antagonism ............................ 146 

5.4.2  SSTR2 antagonism improves counterregulation following recurrent hypoglycemia ................................................................................... 150 

5.4.2.1  SSTR2 antagonism increases glucagon counterregulation. ........... 150 

5.4.2.2  SSTR2 antagonism preserves epinephrine counterregulation. ....... 152 

5.4.2.3  SSTR2 antagonism preserves corticosterone counterregulation. ... 154 

5.4.3  Discussion of other mechanisms for reduced counterregulation following recurrent hypoglycemia ...................................................................... 155 

5.4.4  Why might SSTR2 antagonism not have as great of an effect on recurrent hypoglycemia as acute hypoglycemia? .................................................. 158 

5.5  Study 2 Conclusions .................................................................................... 160 

Chapter 6 Study Summaries and General Discussion ........................................... 177 

6.1  Summary of Study 1 .................................................................................... 178 

6.2  Summary of Study 2 .................................................................................... 179 

6.3  General discussion ...................................................................................... 180 

6.3.1  Limitations of the present study .......................................................... 180 

6.3.2  Hypothesizing a mechanism of action: CNS effects? ............................... 182 

6.3.2.1  Does peripherally administered SSTR2 antagonist enter the brain?185 

6.3.3  Brief evaluation of tachyphylactic effects of SSTR2 antagonism ................ 186 

6.3.4  Potential side effects of SSTR2 antagonism ........................................... 187 

Chapter 7 Future Directions ................................................................................ 190 

7.1  Delineating a mechanism of action for SSTR2 antagonism ................................. 190 

7.2  Testing the efficacy of SSTR2 antagonism in other diabetic models ..................... 191 

7.3  Effect of SSTR2 antagonism to non-hypoglycemia stimuli .................................. 192 

7.4  Interaction between glycemia, insulin, and somatostatin antagonism on glucagon release ...................................................................................................... 192 

Chapter 8 Conclusions ......................................................................................... 194 

8.1  Importance of contributions to current body of knowledge ................................ 194 

8.2  Potential clinical contributions ....................................................................... 195 

Chapter 9 References .......................................................................................... 197 

xi

List of Tables

Table 4-1. Metabolic parameters of rats that underwent hypoglycemia clamps or control

experiments in the absence of insulin.. ..................................................................... 129

Table 4-2. Basal plasma counterregulatory hormone levels for rats that underwent

hypoglycemia clamp and control experiments in the absence of insulin ......................... 130

Table 4-3. Plasma insulin levels during hypoglycemia clamps .................................... 131

Table 4-4. Plasma counterregulatory hormone and insulin levels during control

experiments in the absence of insulin ....................................................................... 132

Table 4-5. Quantitative mRNA expression of liver PEPCK1 and G6Pase following

hypoglycemia clamp (10 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist) and control

experiments in the absence of insulin (0 U/kg, 3000 nmol/kg/h) .................................. 133

Table 4-6. Metabolic parameters of non-diabetic rats treated with SSTR2 antagonist

(N+SSTR2a) that underwent hypoglycemia clamp experiments ................................... 134

Table 4-7. Pancreatic somatostatin protein levels (pmol/mg protein) and plasma

somatostatin levels (pmol/mL) in all hypoglycemic rats .............................................. 135

Table 5-1. Metabolic parameters for rats that underwent recurrent hypoglycemia

treatment. ........................................................................................................... 173

Table 5-2. Body weight and amount of insulin administered via iv bolus and iv infusion,

and total insulin administered in both groups on both days ......................................... 174

Table 5-3. Basal plasma hormone levels on Experimental Day 1 (ExptD1) and

Experimental Day 2 (ExptD2) in rats that underwent SSTR2a-treated and control

hypoglycemia experiments ..................................................................................... 175

Table 5-4. Plasma insulin levels (ng/mL) during ExptD1 and ExptD2 in saline-SSTR2a

treatment and saline-saline control groups ............................................................... 176

xii

List of Figures

Figure 1-1. Linear structures of somatostatin, somatostatin-28, and SSTR2 antagonist

PRL-2903. ............................................................................................................. 53 

Figure 1-2. Structure of SSTR2 antagonist PRL-2903. ............................................... 53 

Figure 3-1. Structure of streptozotocin. .................................................................. 57 

Figure 4-1. Study 1 protocol design. ...................................................................... 115 

Figure 4-2. Schematic representation of A) hypoglycemia clamp experiments and B)

control experiments in the absence of exogenously administered insulin. ...................... 116 

Figure 4-3. Pilot study: Plasma hormone levels during hypoglycemia clamp (10 U/kg

insulin, 1500 nmol/kg/h SSTR2 antagonist). A) Glucagon. B) Corticosterone. .............. 117 

Figure 4-4. Blood glucose levels during hypoglycemia clamp (10 U/kg insulin, 3000

nmol/kg SSTR2 antagonist). ................................................................................... 118 

Figure 4-5. Plasma hormone levels during hypoglycemia clamp (10 U/kg insulin, 3000

nmol/kg/h SSTR2 antagonist). A) Glucagon. B) Corticosterone. C) Glucose infusion. ... 119 

Figure 4-6. Plasma A) epinephrine and B) norepinephrine levels during hypoglycemia

clamp (10 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist) ........................................ 120 

Figure 4-7. Blood glucose levels during hypoglycemia clamp using low insulin dose (5 U/kg

insulin, 3000 nmol/kg SSTR2 antagonist).. ............................................................... 121 

Figure 4-8. Plasma hormone levels during hypoglycemia clamp (5 U/kg insulin, 3000

nmol/kg/h SSTR2 antagonist). A) Glucagon. B) Corticosterone. C) Glucose infusion. ... 122 

Figure 4-9. Plasma A) epinephrine and B) norepinephrine levels during hypoglycemia

clamp (5 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist). ......................................... 123 

xiii

Figure 4-10. Plasma growth hormone levels during hypoglycemia clamp (5 U/kg insulin,

3000 nmol/kg/h SSTR2 antagonist) ......................................................................... 124 

Figure 4-11. Blood glucose levels during control experiments in the absence of insulin (0

U/kg, 3000 nmol/kg/h). ......................................................................................... 125 

Figure 4-12. Pancreatic protein content of A) glucagon and B) somatostatin, and C)

plasma somatostatin, following hypoglycemia clamp (10 U/kg insulin, 3000 nmol/kg/h

SSTR2 antagonist) and control experiments in the absence of insulin (0 U/kg, 3000

nmol/kg/h). ......................................................................................................... 126 

Figure 4-13. Blood glucose levels during hypoglycemia clamp in non-diabetic rats. A) 10

U/kg insulin, 1500 nmol/kg/h SSTR2 antagonist; B) 10 U/kg insulin 3000 nmol/kg/h SSTR2

antagonist; C) 5 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist. ............................... 127 

Figure 4-14. Effect of SSTR2 antagonist on plasma glucagon (A-C) and plasma

corticosterone (D-F) at the various dosing combinations of insulin and SSTR2 antagonist in

non-diabetic rats. .................................................................................................. 128 

Figure 5-1. Study 2 protocol design. ....................................................................... 161 

Figure 5-2. Pilot study: A) Glycemia during 2 consecutive days of hypoglycemia (i.e.

without prior recurrent hypoglycemia). B) Plasma glucagon levels during 2 consecutive

days of hypoglycemia. Insulin administered via s.c. injection. ..................................... 162 

Figure 5-3. Pilot study: A) Glycemia during 2 consecutive days of hypoglycemia (i.e.

without prior recurrent hypoglycemia). B) Plasma glucagon levels during 2 consecutive

days of hypoglycemia. Insulin administered via i.v. bolus + infusion. . ........................ 163 

Figure 5-4. Blood glucose levels during recurrent hypoglycemia treatment (5 episodes

over 3 days). ........................................................................................................ 164 

Figure 5-5. Blood glucose levels during hypoglycemia experiments in saline-SSTR2a

treatment group ................................................................................................... 165 

Figure 5-6. Blood glucose levels during hypoglycemia experiments in saline-saline control

group. ................................................................................................................. 166 

Figure 5-7. Enlarged scale showing blood glucose levels during hypoglycemia experiments

in the A) saline-SSTR2a treatment group and B) saline-control group ........................... 167 

xiv

Figure 5-8. Plasma glucagon during ExptD1 and ExptD2 in A) saline-SSTR2a treatment;

B) saline-SSTR2a treatment with rats that did not experience hypoglycemia omitted; and C)

saline-saline control groups.. .................................................................................. 168 

Figure 5-9. Plasma epinephrine during ExptD1and ExptD2 in A) saline-SSTR2a treatment;

B) saline-SSTR2a treatment with rats that did not experience hypoglycemia omitted; and C)

saline-saline control groups. ................................................................................... 169 

Figure 5-10. Plasma corticosterone during ExptD1and ExptD2 in A) saline-SSTR2a

treatment; B) saline-SSTR2a treatment with rats that did not experience hypoglycemia

omitted; and C) saline-saline control groups ............................................................. 170 

Figure 5-11. Plasma norepinephrine during ExptD1 and ExptD2 in A) saline-SSTR2a

treatment; B) saline-SSTR2a treatment with rats that did not experience hypoglycemia

omitted; and C) saline-saline control groups ............................................................. 171 

Figure 5-12. Comparison of peak incremental hormone responses to acute hypoglycemia

(Study 1) and recurrent hypoglycemia (Study 2) in control and SSTR2 antagonist-treatment

groups. A) Plasma glucagon; B) plasma epinephrine; C) plasma corticosterone.. .......... 172 

xv

List of Abbreviations

ACCORD Action to Control Cardiovascular Risk in Diabetes

ACTH adrenocorticotrophic hormone

ADVANCE Action in Diabetes and Vascular Disease

ANOVA analysis of variance

AP area postrema

ARC arcuate nucleus

AUC area under the curve

AVP arginine vasopressin

BLA basolateral amygdala

BNST bed nucleus of the stria terminalis

CPE carboxypeptidase E

CRH corticotrophin-releasing hormone

DCCT Diabetes Control and Complications Trial

DMH dorsomedial nucleus

DMNV dorsal motor nucleus of the vagus

EDTA ethylenediamine tetraacetic acid

GABA gamma-aminobutyric acid

GE glucose-excited

GHRH growth hormone releasing hormone

GI glucose-inhibited

GPCR G-protein coupled receptor

HAAF hypoglycemia-associated autonomic failure

HPA hypothalamic-pituitary-adrenal

IML intermediolateral column

ip intraperitoneal

iv intravenuous

LC locus coeruleus

LHA lateral hypothalamic area

MPO medial preoptic area

mRNA messenger ribonucleic acid

NOD non-obese diabetic

xvi

NTS nucleus of the solitary tract

PBN parabrachial nucleus

PC1/2 prohormone convertase-1/-2

PCR polymerase chain reaction

PMV portal-mesenteric veins

POMC proopiomelanocortin

PVN paraventricular nucleus

RIA radioimmunoassay

RVLM rostroventrolateral medulla

sc subcutaneous

SCN suprachiasmatic nucleus

SEM standard error of the mean

SON supraoptic nucleus

SSTR1/2/3/4/5 somatostatin receptor type 1/2/3/4/5

STZ streptozotocin

UKPDS United Kingdom Prospective Diabetes Study

VADT Veterans Affairs Diabetes Trial

VMH ventromedial hypothalamus

VMN ventromedial hypothalamic nucleus

1

1

Chapter 1 General Introduction

1 Introduction

1.1 Hypoglycemia

Hypoglycemia is a perturbation to physiological homeostasis and has potentially devastating

consequences. Hypoglycemia is defined as an “abnormally low plasma glucose

concentration that expose[s] the individual to potential harm” by the American Diabetes

Association Workgroup on Hypoglycemia, and quantitatively, it is a measured plasma

glucose level of 3.9 mM (1). Glucose is the predominant fuel to the brain (2). When

arterial plasma glucose levels fall below the physiological euglycemic range (i.e. 3.9 – 6.1

mM), glucose transport to the brain becomes rate limiting to brain glucose metabolism

(3;4). Thus, it is imperative that blood glucose concentrations are maintained above this

level for brain function and ultimately for survival (3;4).

Symptoms arising from iatrogenic hypoglycemia are conventionally classified into two main

categories: i) neuroglycopenic, and ii) neurogenic or autonomic (5). Neuroglycopenic

symptoms stem from glucose deprivation of the brain and include behavioural changes,

fatigue, and cognitive impairments, and confusion (5). Restoration of blood glucose levels

results in recovery from these deficits in brain function (6). In severe neuroglycopenia,

seizures, loss of consciousness, and brain death can ensue, but this is rare and associated

with profound and prolonged hypoglycemia (7). Neurogenic symptoms arise from the

autonomic nervous system reacting to the perception of hypoglycemia. Adrenergic

2

neurogenic symptoms mediated by catecholamines include shakiness/tremor, a pounding

heart, and nervousness/anxiety, while cholinergic neurogenic symptoms mediated by

acetylcholine include sweatiness, hunger, and a tingling sensation (8). In healthy

individuals, these symptoms of hypoglycemia often do not present until plasma glucose

levels reach a threshold of approximately 3.0 mM (9-11). Neurogenic symptoms are

present (3.2 mM) before neuroglycopenic symptoms (2.8 mM) and cognitive impairment

(2.7 mM) (12).

1.1.1 Normal sensing of hypoglycemia: sites of glucose detection

Hypoglycemia is a fairly uncommon event except in individuals that use glucose-lowering

medications, particularly insulin and sulphonylureas or other insulin secretagogues to treat

diabetes, or in patients with insulinomas (11). This is because the body has several

physiological defense mechanisms to counter decreases in blood glucose to restore blood

glucose from hypoglycemia to euglycemia. In healthy subjects, hypoglycemia elicits a

characteristic succession of responses. Glucose sensors are responsible for the detection of

glycemic decline and initiate an orchestrated series of responses mediated via neural

networks that integrate the various afferent inputs and instigate efferent responses,

particularly as they relate to hypoglycemia. Glucose sensors important for glucose

counterregulation are found in three primary areas: the hepatic portal-mesenteric veins

(PMV), the hypothalamus, and the brainstem (13;14).

The portal vein can also sense a decrease in plasma glucose via a GLUT2-dependent glucose

sensing mechanism and relays this signal to the central nervous system via a vagus nerve

afferent (15-18). It has been demonstrated that normalizing glucose levels in the PMV can

suppress hypoglycemia-induced food intake (19) and sympathoadrenal responses (20;21).

3

Hepatoportal glucose sensors also stimulate glucose storage in the liver, muscle, and

adipose tissue (22).

In the hypothalamus, established key areas involved in glucose detection include the

ventromedial hypothalamus (VMH) (specifically the ventromedial hypothalamic nuclei (VMN)

and arcuate nuclei (ARC)) (23-26), lateral hypothalamic area (LHA) (27-29), and

paraventricular nuclei (PVN) (30-32) in which glucose sensing neurons have been located.

The VMH integrates forebrain neuronal input with ascending information from the brainstem

and then sends output to regions involved in control of various behaviours and physiologic

responses (33). These hypothalamic areas identified for glucose sensing also have critical

roles for glucagon and catecholamine counterregulation in response to hypoglycemia (34-

40), which will be discussed below.

In the brainstem, glucose sensing neurons are abundant in the nucleus of the solitary tract

(NTS) (15;41-43), area postrema (AP) (44), and dorsal motor nucleus of the vagus (DMNV)

(45). These glucose sensing regions of the brainstem have been implicated in the

regulation of glucose homeostasis (46-48).

More recently, nuclei in the medial amygdala have also been implicated as an important

glucose sensing area that also elicits counterregulatory responses (49). Detailed

mechanisms involved with sensing hypoglycemia and neuronal integration have been

recently reviewed (13;14;18). In summary, perturbation of glucose homeostasis triggers

central integration of the various glucose-detection signals, and an autonomic response is

initiated that activates sympathetic (neural and adrenomedullary) and parasympathetic

outflow (50). Taken together, these counterregulatory responses involve inhibition of

4

endogenous insulin secretion and activation of several hormonal and neuroendocrine

systems to restore blood glucose to euglycemia.

1.1.2 Normal counterregulatory hormone responses to hypoglycemia

1.1.2.1 Insulin.

First and foremost, with declining blood glucose levels, insulin secretion must cease

(51;52), and this alleviates both its suppressive effect on hepatic glucose production and its

stimulatory effect of glucose utilization in muscle and adipose tissue. Reduced insulin

secretion also lowers insulin-mediated inhibition of lipolysis and protein catabolism and

stimulation of lipogenesis and protein anabolism, which results in increased plasma levels of

free fatty acids, glycerol, and amino acids, the products of which can serve as substrates or

energy sources for gluconeogenesis (53;54). Reduced insulin secretion is believed to be

due to a direct effect of hypoglycemia on pancreatic -cells (51;55). Typically, the decrease

in pancreatic insulin secretion occurs when the blood glucose threshold reaches 4.6 mM (3).

Sympathoadrenal activation in the form of norepinephrine from pancreatic sympathetic

nerves and epinephrine from the adrenal medulla may also play a role in inhibiting insulin

secretion via 2-adrenergic stimulation (56). It should be noted that the catecholamines

can also stimulate insulin via 2-adrenergic receptors, but the 2-adrenergic effect appears

to predominant, at least in human islets (57)

1.1.2.2 Glucagon.

In addition to the decrement in insulin, several other counterregulatory hormones work

together to normalize blood glucose levels in response to hypoglycemia. It is well known

that glucagon is key in restoring euglycemia during acute hypoglycemia (58;59). Glucose

recovery from hypoglycemia is impaired by approximately 40% when glucagon secretion is

inhibited by somatostatin infusion (60). This dependence on the glucagon response for

5

hypoglycemic recovery is true for both short-term (59-61) and prolonged (62) episodes of

hypoglycemia. Glucagon helps to restore euglycemia by stimulating hepatic glucose

production via glycogenolysis, or it favours gluconeogenesis when precursors (i.e. lactate,

amino acids, and glycerol) are abundant (63).

Increments in glucagon secretion can result from direct effects of hypoglycemia on

pancreatic -cells (64). The glycemic threshold of approximately 3.8 mM triggers glucagon

release (9;10). Other mechanisms of the secretory glucagon response to hypoglycemia

include increased autonomic inputs (i.e. sympathetic neural, parasympathetic neural, and

adrenomedullary) and decreased intraislet insulin (50). During hypoglycemia, the

sympathetic (65;66) and parasympathetic (67;68) nerves and the adrenal medulla are

activated – releasing norepinephrine (69;70), acetylcholine (71), and epinephrine (72;73),

respectively – all of which result in increased glucagon release. Catecholamines stimulate

-cell glucagon secretion via 2- and 2-adrenergic receptors (69;74;75), whereas

acetylcholine acts via M1 and M3 muscarinic receptors (76;77). Decreased intraislet insulin

secretion during hypoglycemia is believed to be one of the signals involved in glucagon

secretion (78-80). More recent work has suggested that a decrement in zinc, a co-secretory

product of insulin release, and not necessarily insulin itself, provides the trigger to initiate

glucagon secretion during hypoglycemia since zinc acts on KATP-channels to hyperpolarize -

cells (81;82), but this view has been challenged (83). -aminobutyric acid (GABA), which is

co-localized in -cells with insulin, also suppresses glucagon release (84). Functional GABAA

receptors are expressed on -cells, and activation of these receptors by GABA secreted by

-cells mediates chloride ion influx thereby hyperpolarizing the -cell and inhibiting

glucagon release (85-87). Intraislet somatostatin exerts a tonic suppressive effect on

glucagon release (88;89). It was first postulated nearly 20 years ago that relative

6

abundance of pancreatic somatostatin in diabetic dogs may contribute to the poor glucagon

response to hypoglycemia (90), and this concept will be discussed in detail throughout this

thesis. Ghrelin is another peptide predominantly produced by the stomach but which is also

synthesized in small amounts in the pancreatic islet. The ghrelin receptor is widely

expressed in the adult rat islet and has been co-localized with glucagon-positive cells (91).

Locally released ghrelin could have a paracrine effect on -cell glucagon release, perhaps

promoting glucagon release during hypoglycemia. However, studies in perfused rat

pancreas indicate that concentrations of ghrelin within the physiological range (0.5–3 nM)

had no effect on basal or arginine-stimulated glucagon release, despite mildly inhibiting

both stimulated insulin and somatostatin release (92). It has also been postulated that the

-cell may sense hypoglycemia directly and secrete glucagon in response to low glucose

concentrations as has been observed in the perfused isolated pancreas (93). However,

lowering glucose levels in isolated -cells failed to stimulate glucagon secretion (94), which

suggested no direct role of hypoglycemia on stimulating glucagon release. More recent

evidence suggests that the direct stimulatory effect of low glucose on glucagon secretion by

the -cell may be of less significance as compared to the stimulatory influences of other

factors (95;96). This is supported by the intraislet insulin hypothesis which involves a

“switch-off” concept in which the -cell reduces insulin secretion as a key mechanism to

initiate the glucagon response to hypoglycemia (80;97). This concept as it pertains to

diabetes will be discussed in section 1.4.1.

In addition to these factors, central nervous system regulation of glucagon release has also

been demonstrated. Lesioning of the VMH caused 50-60% and 75-80% decreases in

glucagon and catecholamine responses during mild (3.0 mM) and severe (2.5 mM)

hypoglycemia, respectively (34). Glucagon secretion can be induced with local

7

glucoprivation in the VMH by direct 2-deoxyglucose injection (35), and hypoglycemia-

induced glucagon secretion can be suppressed by direct VMH injection of glucose (36).

Furthermore, glutamate release by neurons of the VMH has also been implicated as an

important mechanism in glucagon counterregulation to hypoglycemia (98). The VMH has

been shown to be part of the neurocircuitry that links sympathetic and parasympathetic

innervation of the pancreas to regulate glucagon release, and Buijs et al. reports other key

brain centres including the suprachiasmatic nucleus (SCN), ARC, dorsomedial nucleus

(DMH), locus coeruleus (LC), NTS, and DMNV and intermediolateral column (IML) of the

spinal cord to be involved in this neurocircuitry (99).

1.1.2.3 Epinephrine.

Epinephrine has an important role in augmenting blood glucose levels, although the

epinephrine response may not be critical in correcting hypoglycemia unless glucagon

secretion is deficient (59;100;101), which often occurs in type 1 diabetes or long-standing

type 2 diabetes. This will be further discussed below. It has been suggested that in type 1

diabetes, a robust epinephrine response can compensate for an absent glucagon response

(102). Hypoglycemia is one of the most potent stimulators of epinephrine release from the

adrenal medulla (103) and has been demonstrated to elevate plasma epinephrine levels by

more than 60-fold (103;104). Hypoglycemia activates sympathetic nervous activity in

muscle (105;106) and the pancreas (65;67). The release of epinephrine is triggered when

arterialized venous blood glucose levels fall to approximately 3.8 mM, which is similar to the

glycemic threshold for glucagon secretion (9;10). Epinephrine both stimulates glucose

production as well as limits glucose utilization directly by inhibiting glucose uptake by

peripheral tissues and indirectly by suppressing insulin release (107). Epinephrine

stimulates hepatic glycogenolysis via 1- and 2-adrenergic receptors (108). Epinephrine

8

stimulates hepatic, and to some extent renal, gluconeogenesis, also by mobilizing

gluconeogenic substrates. The effect of epinephrine to reduce glucose uptake involves

limiting glucose utilization by insulin-sensitive tissues via direct effects and also by

increasing non-esterified fatting acids levels. Catecholamine release from the adrenal gland

can also stimulate synthesis of glucocorticoids (109).

1.1.2.4 Norepinephrine.

Although some norepinephrine is also released by the adrenal medulla (approximately 20%

of its total catecholamine release is norepinephrine), the primary source of circulating

norepinephrine is from sympathetic nerves innervating blood vessels. Normally, most of the

norepinephrine released by sympathetic nerves is taken back up by the nerves (some is also

taken up by extra-neuronal tissues) where it is metabolized. Thus, only a small amount of

norepinephrine enters the blood into the peripheral circulation. However, at times of high

sympathetic nerve activation, the amount of norepinephrine entering the blood increases

dramatically (108). Norepinephrine has also been shown to increase blood glucose levels

during hypoglycemia via mechanisms similar to those of epinephrine (110). Yet during

hypoglycemia, plasma norepinephrine levels are lower compared with epinephrine (i.e.

approximately 5-fold as compared to 60-fold elevations of epinephrine) (103;111). During

hypoglycemia, norepinephrine is released not only from sympathetic nerve terminals, but

also in smaller quantities from the adrenal medulla (112-114). Increases in norepinephrine

within muscle and adipose tissue are observed in response to hypoglycemia implicating

activation of sympathetic nerves in these glucoregulatory tissues. In muscle,

norepinephrine and epinephrine can stimulate glycogenolysis and release lactate via a 2-

adrenal mechanism and decrease insulin-stimulated glycogen synthesis (108). In adipose

9

tissue, norepinephrine and epinephrine mobilize free fatty acids by stimulating lipolysis also

mediated through -adrenergic receptors (115;116).

1.1.2.5 Growth hormone and glucocorticoids.

Suppression of insulin secretion and activation of glucagon and epinephrine secretions are

the primary defenses to safeguard against hypoglycemia. Growth hormone and

glucocorticoids also help to promote the restoration of euglycemia during prolonged

hypoglycemia (i.e. ≥ 3 hours) (64;117;118), whereas their effects during short term

hypoglycemia are not as strong. Growth hormone and glucocorticoid responses are

triggered at glycemic thresholds of approximately 3.7 and 3.2 mM, respectively (9;10).

Growth hormone and glucocorticoids – cortisol in humans and corticosterone in rodents –

act by decreasing glucose utilization and supporting glucose production (117;118). During

prolonged hypoglycemia, the lack of either growth hormone or glucocorticoid

counterregulation results in a hypoglycemia of greater severity which even enhanced

glucagon or epinephrine counterregulation cannot overcome (117;118). Growth hormone

secretion from the anterior pituitary is primarily regulated by growth hormone release

hormone (GHRH) and somatostatin release from the hypothalamus. Insulin-induced

hypoglycemia, L-dopa administration, and arginine infusion indirectly stimulate growth

hormone secretion, and one study has suggested that unlike the other stimuli,

hypoglycemia can elicit increases in growth hormone secretion independently of GHRH and

somatostatin regulation (119). During hypoglycemia, an increase in growth hormone is

involved in enhancing glucose production, limiting glucose utilization, and stimulating

lipolysis to increase circulating plasma free fatty acids, glycerol, and -hydroxybutyrate,

which can provide gluconeogenic substrates and well as fuels to spare glucose (117).

10

In response to hypoglycemia, the hypothalamic-pituitary-adrenal (HPA) axis is activated,

and this results in the release of cortisol/corticosterone from the adrenal cortex (120).

Activation of the HPA axis involves the release of corticotrophin-releasing hormone (CRH)

and arginine vasopressin (AVP) from neurons of the paraventricular nucleus of the

hypothalamus into the median eminence. Here, these neuropeptides travel via the

hypophyseal portal circulation to the anterior pituitary where they stimulate the production

of adrenocorticotrophic hormone (ACTH) from its precursor, proopiomelanocortin (POMC).

ACTH is released into the peripheral circulation where is acts on the adrenal cortex to

release cortisol/corticosterone, which ultimately inhibit further HPA activity (121;122). This

negative feedback control is mediated by cortisol/corticosterone acting on glucocorticoid

receptors (GRs) in the hippocampus, PVN, and anterior pituitary (for suppressing the stress

response) or on mineralocorticoid receptors at the hippocampus (for tonic basal inhibition of

the HPA axis) (123). During hypoglycemia, glucocorticoids act on glucocorticoid receptors

on the liver to stimulate glucose production directly by enhancing gluconeogenic enzymes

and indirectly by increasing the pool of readily available substrates (124). Glucocorticoids

reduce insulin-mediated glucose uptake and utilization by muscle and adipose tissue

(125;126). In support of the primary counterregulatory defenses, glucocorticoids can

potentiate the effects of glucagon and epinephrine on the liver and suppress insulin’s

inhibitory effect of glucose production (127;128).

1.1.2.6 Glucose autoregulation.

Glucose autoregulation is endogenous glucose production independent of the other

aforementioned hormones and neural glucoregulatory mechanisms, and it is inversely

proportional to ambient plasma glucose levels (50). This has been observed in humans

during marked hypoglycemia (110;129). It is only at very severe hypoglycemia (i.e. 1.7 –

11

1.9 mM) that glucose autoregulation may occur (129;130). It is believed that glucose

autoregulation plays a relatively minor role in glucose counterregulation (3).

1.2 Hypoglycemia in diabetes

1.2.1 Clinical Significance – Hypoglycemia in type 1 diabetes mellitus

Hypoglycemia is rare in people without diabetes (11;63;131-133). However, hypoglycemia

is one of the most serious acute complications of well-controlled type 1 diabetic patients and

often represents the limiting factor to insulin therapy (134-140). Based on conservative

estimates, 10% of type 1 diabetic individuals using conventional insulin therapy and 25% of

diabetic individuals using intensive insulin therapy experience at least one bout of severe,

debilitating hypoglycemia, often with seizure or coma, in a given year (138;139;141).

Furthermore, patients practicing conventional insulin therapy undergo an average of one

bout of mild to moderate symptomatic hypoglycemia per week while those being intensively

treated suffer from two such episodes weekly (142). It can thus be projected that over 40

years of type 1 diabetes, an average patient may experience 2000-4000 bouts of

symptomatic hypoglycemia (143). Hence, iatrogenic hypoglycemia is evidently problematic

for type 1 diabetic patients attempting to manage their plasma glucose levels in a range

commonly recommended by practitioners to curtail diabetic complications resulting from

poor glycemic control. To further complicate matters, the strongest predictor of severe

hypoglycemia was a history of severe hypoglycemia (144). This suggests any bout of

hypoglycemia can predispose an individual to suffer subsequent bouts of hypoglycemia.

The topic of recurrent hypoglycemia will be discussed below.

12

1.2.2 Clinical Significance – Hypoglycemia in type 2 diabetes mellitus

Individuals with type 2 diabetes generally experience less frequent severe hypoglycemia

than those with type 1 diabetes as has been reported by two separate clinical trials (143).

Individuals with type 2 diabetes are at a two- to four-fold greater risk of serious

cardiovascular outcomes compared with those without diabetes (145). As with type 1

diabetes, the ultimate goal of well-controlled glycemia to reduce the risk of diabetic

complications is still elusive due to an increased threat of hypoglycemia that accompanies

intensive treatment (146). Treatment-induced hypoglycemia is a limiting factor of glycemic

management not only in individuals with type 1 diabetes, but also in many with advanced

type 2 diabetic individuals in which there is an absolute endogenous insulin deficiency

(63;147-149). Overall, hypoglycemia is less frequent in type 2 diabetes than in type 1

diabetes, but hypoglycemia becomes progressively more frequent and limiting to glycemic

control in advanced type 2 diabetes (4). Patients with type 2 diabetes recently started on

insulin had rates of hypoglycemia which were similar to those patients who were taking

sulphonylureas (150). However, individuals with type 2 diabetes treated with insulin for

over 5 years and whose endogenous insulin production as measured by stimulated C-

peptide was low had significantly higher rates of hypoglycaemia than individuals with

shorter duration of insulin treatment or individuals treated with sulphonylureas, including

severe episodes (150;151). Strict glycemic control remains the most effective strategy to

reduce the development and/or progression of microvascular disease. Interestingly,

however, recent clinical trials in type 2 diabetic patients – Action to Control Cardiovascular

Risk in Diabetes Study Group (ACCORD) (152), Action in Diabetes and Vascular Disease

(ADVANCE) (153), and Veterans Affairs Diabetes Trial (VADT) (154) – reported that

reducing HbA1C to below 7% did not significantly reduce cardiovascular mortality.

However, it has been suggested that the study duration of these aforementioned trials may

13

have been too short to show a significant effect of intensive glycemic control on

macrovascular complications (146). The UKPDS study, on the other hand, showed

significantly lower relative risk reduction for myocardial infarctions and overall mortality in

patients that receive intensive treatment compared with those who received conventional

treatment in the 10-year follow-up study (155). Thus, the effect of intensive control of

glycemia to reduce cardiovascular morbidity has been demonstrated, but it may be limited

to younger patients with shorter duration of diabetes and no prior cardiovascular disease

(156).

Despite the benefits of well-controlled glycemia on cardiovascular morbidity, intensive

treatment to control glucose is associated with a significantly greater incidence of severe

hypoglycemic events (152-154). Mild hypoglycemia events were also more frequent in the

intensive control arm as compared to conventional control arm in the ADVANCE trial (153).

In the ACCORD trial, there was a 3-fold greater incidence of severe hypoglycemia in the

intensive treatment group as compared with conventional treatment group (152). The

VADT study reported 3-fold greater incidence of hypoglycemia with symptoms, and a 5-fold

greater incidence of hypoglycemia without symptoms, in the intensive versus standard

therapy arms (154). This is problematic since in the VADT, severe hypoglycemia was the

most powerful predictor of cardiovascular death. Likewise in ACCORD, patients with severe

hypoglycemia had higher mortality than those without in both the intensive and

conventional treatment study arms. Hypoglycemia is associated with activation of

adrenergic responses and prolonged QTc intervals and cardiac arrhythmias (157), which can

contribute to increased cardiovascular mortality. In addition, diabetic autonomic

neuropathy may also render patients more vulnerable to adverse cardiovascular outcomes

(158). Thus, intensive treatment to keep HbA1C below 7% reduces the risk of

14

macrovascular complications in individuals with type 2 diabetes, but the corollary is that

intensive glucose control is associated with frequent episodes of severe hypoglycemia and

consequent cardiovascular events leading to mortality.

1.3 Defective counterregulation in diabetes

Normal counterregulatory responses to hypoglycemia have been described above as it

applies to healthy individuals. However, hypoglycemia in type 1 diabetic patients is the

consequence of an interplay between insulin excess as well as compromised glucose

counterregulation (159). In type 2 diabetes, both a history of previous hypoglycemia and

duration of insulin treatment are significant predictors of hypoglycemia (148;160). As

patients progress with type 2 diabetes, they approach the insulin-deficient end of this

disease until they reach the stage of being absolutely endogenous insulin deficient. Then

patients are thus similar to individuals with type 1 diabetes in that compromised glucose

counterregulation may include defective hormone responses to restore euglycemia,

hypoglycemia unawareness, and elevated glycemic thresholds (i.e. lower glycemia required)

for responses to hypoglycemia (161).

1.4 Mechanisms of defective counterregulation in diabetes

Fundamental alterations in glucose counterregulation occur in individuals with established

type 1 and type 2 diabetes (159;162). When diabetes is fully developed, endogenous

insulin secretion in completely absent (163;164), and plasma insulin concentrations reflect

that those which are provided by exogenously administered insulin and do not diminish

when blood glucose levels fall. Thus the first defense against hypoglycemia, the decrement

15

of endogenous insulin secretion, is non-existent (159;162). As patients with type 2

diabetes approach the insulin-deficient end of this disease, they become similar to

individuals with type 1 diabetes in that both have absolute -cell failure, which causes the

loss of the insulin decrement in response to hypoglycemia (4).

1.4.1 Glucagon counterregulation in impaired in diabetes

The glucagon response, the second defense to combat hypoglycemia, is also lost in

established type 1 diabetes (165;166) and in advanced type 2 diabetes (167;168). In type

1 diabetes, this usually occurs within the first few years of onset of the disease (165;166).

This defect may be specific to insulin-induced hypoglycemia as a stressor since glucagon

counterregulation to other stressors, such as arginine infusion or intracerebroventricular

carbachol infusion, in diabetic subjects or animals are still intact (166;169;170). Although it

has been shown that the impairment is linked to a loss of residual -cell function (171), the

exact mechanisms underlying the defect in glucagon response still remain unclear.

Hypotheses related to impaired glucagon counterregulation as a result of loss of -cell

function include a requirement of a decrement of intraislet insulin release (172;173) as well

as a -cell insulin “switch-off” hypothesis (82;97) to trigger glucagon secretion in response

to hypoglycemia. Other hypotheses include chronic hyperglycemia since it was

demonstrated that restoration of normal glycemia independent of insulin can improve

glucagon counterregulation (174), increased -cell sensitivity to exogenous insulin as a

result of the absence of -cell function (175), and a defect in autonomic response to

hypoglycemia (175), and in particular defective sympathetic innervation of the islet to

release glucagon in response to hypoglycemia (176). Reduced autonomic responses to

hypoglycemia may be the result of up-regulated brain glucose transport since it has been

demonstrated that this is a prominent mechanism after long repeated exposure to

16

hypoglycemia (177;178). In -cell-deficient STZ-diabetic rats treated with insulin, it was

demonstrated that the glucagon response to insulin-induced hypoglycemia can be enhanced

by pancreatic infusion and subsequent switch-off of somatostatin (at hypoglycemia),

thereby supporting the notion that glucagon counterregulation is a general disinhibition

phenomenon and can be enhanced by switch-off of -cell-suppressing intrapancreatic

signals other than insulin or its co-secretory products (179).

In type 2 diabetes, which tends to be a more heterogeneous disease than type 1 diabetes,

investigations of how hypoglycemic counterregulation is altered in this disease must also

consider factors such as age, duration of diabetes, -cell reserve, type of therapy (i.e. diet,

exercise, oral medications, insulin, etc.), body fat composition, co-morbidities, and

metabolic control (180;181). The glucagon response to hypoglycemia in type 2 diabetes has

been shown to be preserved in some studies (182-185), but defective in other reports

(167;168;186-188). An absolute insulin deficiency associated with advanced type 2

diabetes is suggested to underlie the glucagon impairment in advanced type 2 diabetes

(4;161). A study by Segel et al. separated the glucagon responses of type 2 diabetic

subjects on insulin therapy, who represent an insulin-deficient group, from type 2 diabetic

subjects on oral medications, who represent the insulin-sufficient group, and demonstrates

that insulin-deficient individuals, but not those with endogenous insulin, have diminished

glucagon counterregulation to hypoglycemia (168).

Besides these hypotheses of local factors responsible for impaired glucagon

counterregulation, several mechanisms related to the central nervous system have also

been posited. Local glucopenia in the absence of peripheral hypoglycemia induced by 2-

deoxyglucose injection into the VMH stimulates glucagon and epinephrine responses in non-

diabetic rats, but these responses were markedly suppressed in diabetic rats (189). It is

17

speculated that the impairment of the glucagon response to hypoglycemia in diabetes in

which there is an absolute insulin deficiency might result in part from the simultaneous

increase in insulin at the levels of the VMH and in the pancreatic islet as a result of

exogenous insulin administration (190). Recently, it was demonstrated that insulin

microinjected into the VMH can suppress the glucagon response to hypoglycemia induced by

phloridzin (i.e. hypoglycemia not induced by insulin) by approximately 40% and that VMH

blockade of insulin via insulin immunoneutralization can amplify the glucagon response by

more than 2-fold (190). However, this has yet to be demonstrated in a diabetic model. The

role of glucose in the VMH has also been implicated as mechanism of influencing

hypoglycemic counterregulation. Local delivery of glucose into the VMH suppresses

glucagon and epinephrine responses to hypoglycemia (36;191). When glucose levels

increase in the VMH, an increased in the ATP:ADP ratio in local glucose-excited neurons

leads to closure of KATP channels, membrane depolarization, Ca2+ influx via voltage-

dependent calcium channels, and release of neurotransmitters such as GABA (39;192;193).

GABA acting in the VMH blunts glucagon and epinephrine counterregulation to

hypoglycemia, whereas inhibiting GABA actions can stimulate these counterregulatory

responses (39). A recent study provided the link between the effects of VMH glucose and

GABA by demonstrating the glucose administrated in the VMH dose-dependently activates

local GABA release, which consequently dose-dependently suppresses glucagon and

epinephrine responses to hypoglycemia (191). Studies have also demonstrated a role for a

different population of glucose-sensing neurons of the VMH, the glucose-inhibited (GI)

neurons, in glucagon counterregulation. GI neurons increase their activity in response to

low glucose levels (194). AMP-activated protein kinase (AMPK) acts as an intracellular fuel

gauge which responds to a rise in the intracellular AMP:ATP ratio and is thus implicated in

the glucose sensing pathway used by GI neurons (195). Activation of AMPK in the VMH with

18

locally administered 5-aminoimidazole-4-carboxamide (AICAR) lowered the exogenous

glucose infusion rate and increased glucose production in response to hypoglycemia in non-

diabetic rats (196) and amplified the glucagon responses to hypoglycemia in diabetic rats

(197).

1.4.2 Epinephrine counterregulation in diabetes

The epinephrine response to hypoglycemia is critical in a setting of deficient glucagon

counterregulation to hypoglycemia (4). Individuals lacking both glucagon and epinephrine

counterregulation are at a 25-fold or greater risk of severe hypoglycemia compared to

individuals lacking glucagon but have an intact epinephrine response (198;199). A

combined defect in both glucagon and epinephrine responses is associated with impaired

glucose production in the liver and kidney (143;200). Epinephrine counterregulation to

hypoglycemia has been shown to be both impaired (165;201-205) or not impaired (206-

208) in type 1 diabetes. In non-diabetic humans in which two different doses of insulin

were used to induce hypoglycemia of equivalent depth, it was shown that the higher insulin

dose resulted in a markedly greater increase in epinephrine release than the lower dose

(209). However, this stimulatory effect of high insulin dose did not amplify epinephrine

counterregulation in type 1 diabetic humans, suggesting some adrenomedullary deficit with

insulin-deficient diabetes (210). It has been shown that the glycemic threshold for

epinephrine release is altered such that lower glycemia is needed to elicit a response

(203;211). In type 2 diabetic subjects, the epinephrine response has mostly been reported

to be normal (185;188;212). Interestingly, some have also reported an increased in

epinephrine counterregulation to hypoglycemia in type 2 diabetic subjects (186;213). This

enhanced epinephrine response to hypoglycemia in type 2 diabetes may partially offset

impaired glucagon secretion and counteract the effects of hyperinsulinemia on liver, fat, and

19

skeletal muscle (186) and/or contribute to the promotion of hypoglycemic recovery since an

increase in renal glucose release was also observed in this group (213). The factor of age

should also be considered since it has been reported in non-diabetic individuals that the

glucose threshold for epinephrine and glucagon responses in older patients (2.8 mM) was

significantly lower than in younger patients where the threshold was 3.3 mM (214). Indeed,

although epinephrine and cortisol responses to hypoglycemia in older type 2 diabetic adults

were increased, glucagon and growth hormone responses were reduced when compared to

age-matched non-diabetic controls (215). Furthermore, epinephrine defect in diabetes may

be stressor-specific: although epinephrine was reduced in response to hypoglycemia and

cold stress (216), it was not impaired in response to exercise (204;217). It has also been

shown that poorly controlled diabetic individuals may not suffer from impaired epinephrine

to the same extent of intensively treated diabetic patients (206;207;211;218), suggesting

that prior glycemic control of diabetes may also affect epinephrine counterregulation. In

type 2 diabetic patients, for example, intensive treatment to better control glycemia in has

been shown to blunt epinephrine counterregulation to hypoglycemia (212;219). However,

since tight control of blood glucose levels is also associated with increased frequency of

hypoglycemic episodes, it is also possible that recurrent hypoglycemia leads to the impaired

epinephrine response observed in diabetes, as has been previously observed (220).

Mechanisms related to the regulation of epinephrine counterregulation by the brain, in

particular by the VMH, have been discussed above as it pertains to glucagon. Many of these

studies demonstrate simultaneous influences on both glucagon and epinephrine release in

both non-diabetic (36;39) and diabetic rats (189), suggesting that the counterregulatory

release of both these glucoregulatory hormones are regulated by similar neural pathways

integrated by the VMH. Interestingly, VMH administration of insulin did not affect

20

epinephrine counterregulation (190), suggesting that different neural pathways involving

the VMH also exist. Furthermore, it was demonstrated that inhibition of VMH nitric oxide

(NO) signalling impairs epinephrine, but not glucagon, counterregulation and results in a

greater exogenous glucose infusion rate to prevent hypoglycemia (221). VMH GI neurons

respond to decreased glucose by activating AMPK and stimulating the production of NO

(195;222), and it has been hypothesized that VMH NO production is necessary for

generation of hypoglycemic counterregulation (221).

1.4.3 Norepinephrine, growth hormone, and glucocorticoid counterregulation in diabetes

With respect to the other counterregulatory hormones, it is controversial as to

whether or not neuroendocrine responses are impaired in type 1 diabetes. Studies

examining norepinephrine counterregulation are varied as some studies find decreased

responses (205;223;224) while others do not (202-204;206;225-227). Growth hormone

responses to hypoglycemia have also been reported to be decreased (204;224;228;229) or

unchanged (203;225-227) in type 1 diabetes. Growth hormone responses to hypoglycemia

in type 1 (230) and type 2 (215) diabetic individuals were reduced when compared to age-

matched non-diabetic controls. Lastly, glucocorticoid counterregulation has also been

reported to be decreased (202;205;206;224;226;228) or unchanged (203;204;225;227).

Interestingly, it has been reported that scrupulous avoidance of hypoglycemia could reverse

the impairments of some counterregulatory hormone responses (224). Similar to

epinephrine, prior strict glycemic control, and consequently repeated hypoglycemia, may

also affect the impairment of these responses (206;207;231).

21

1.4.4 Hypoglycemia unawareness in diabetes

Part of the defect in glucose counterregulation to hypoglycemia in diabetes is the clinical

syndrome of hypoglycemia unawareness, which is the impaired awareness of hypoglycemia

(4). More specifically, it is the inability to recognize the neurogenic warning symptoms of

hypoglycemia or the failure for symptoms to occur before the start of neuroglycopenia

(232). With this absence of awareness, a subject developing hypoglycemia is averted from

undertaking protective action to prevent or correct impending hypoglycemia, such as

nutrient ingestion. Hypoglycemia unawareness is associated with a 6-fold increased risk of

severe hypoglycemia (233). It is suggested that the pathophysiology of glucose

counterregulation is similar in type 1 and advanced type 2 diabetes although the timeframe

of these impairments may be different (4). The failure of -cell function is more rapid in

type 1 diabetes but occurs more slowly in type 2 diabetes, and thus the syndromes of

defective glucose counterregulation occur earlier in the former and later in the latter (4).

The attenuation of the sympathoadrenal (mainly sympathetic neural) response causes

hypoglycemia unawareness (233), but several factors are believed to contribute to this

impairment (234). Hypoglycemia unawareness can result from diminished autonomic

responsiveness to hypoglycemia since patients with type 1 diabetes were reported to have

reduced -adrenergic sensitivity, which may play a role to their inability to recognize

adrenergic warning signs to hypoglycemia (235). The lack of the catecholamine response to

hypoglycemia has been attributed to hypoglycemia unawareness since subjects with this

hormone impairment also were asymptomatic for hypoglycemia (236). Elderly patients are

more susceptible to hypoglycemia unawareness than middle-aged patients with type 2

diabetes, and this was found to be independent of differences in hormone counterregulation

between both groups (237). However, the younger cohort had better autonomic and

22

neuroglycopenic symptom scores and were better able to estimate their glycemia to be low

during hypoglycemia. Prior and repeated hypoglycemia also leads to hypoglycemia

unawareness, and it was demonstrated that careful avoidance of hypoglycemia can reverse

hypoglycemia unawareness and improve the epinephrine response (238-240). This may be

associated with improved -adrenergic activity as a result of strict avoidance of

hypoglycemia as has been observed in type 1 diabetic subjects (241).

Hypoglycemia, even if asymptomatic, causes a vicious cycle of recurrent hypoglycemia by

causing hypoglycemia-associated autonomic failure, the clinical syndromes of defective

glucose counterregulation and hypoglycemia unawareness (4), as described below.

1.4.5 Hypoglycemia-associated autonomic failure (HAAF) in diabetes

Hypoglycemia-associated autonomic failure, or HAAF, in diabetes is a concept developed by

Cryer based on the well-established theories that recent antecedent hypoglycemia

(150;153;242), as well as sleep (56) and antecedent exercise (243), cause reduced

sympathoadrenal responses to hypoglycemia, which in turn leads to reduced sympathetic

neural responses and hypoglycemia unawareness as well as defective glucose

counterregulation. Taken together, these impairments lead to a vicious cycle of recurrent

hypoglycemia (63). This concept was originally developed to explain the increased

susceptibility of hypoglycemia in patients with type 1 diabetes (244) and has since also

been applied to explain the increased development of hypoglycemia in advanced type 2

diabetic individuals (168). In type 2 diabetic subjects, recent and repeated hypoglycemia

shifts the glycemic thresholds for autonomic (including adrenomedullary epinephrine) and

symptomatic responses to subsequent hypoglycemia to lower plasma glucose

concentrations, thus contributing to HAAF in type 2 diabetes (168). The concept of HAAF is

23

widely accepted and quoted by many others with respect to both type 1 and type 2 diabetes

(181;219;245-247).

1.5 Recurrent hypoglycemia in non-diabetic subjects

Recurrent hypoglycemia has been shown to blunt subsequent counterregulation to

hypoglycemia in healthy, non-diabetic subjects (248-251). For example, it has been

consistently demonstrated that glucagon and epinephrine responses to hypoglycemia are

diminished after recurrent hypoglycemia in non-diabetic humans (248;250-254) and rats

(255). Prior bouts of hypoglycemia blunted epinephrine, but not norepinephrine, responses

(111;256). Prior bouts also impaired cognitive function during subsequent hypoglycemia in

non-diabetic rats (254;257). Whether impairments exist in the other neuroendocrine

counterregulation, however, is debatable. Norepinephrine (248;251) and cortisol (248;250)

responses have been reported to be reduced in some studies, but these findings have been

inconsistent.

1.6 Recurrent hypoglycemia in diabetes

Recurrent hypoglycemia increases the susceptibility of individuals to subsequent bouts of

hypoglycemia since it contributes to both defective hormone counterregulation and reduced

symptom recognition (hypoglycemia unawareness) (4). HAAF, as explained above, is

relevant in both type 1 diabetes (203) and type 2 diabetes of long duration in which there is

an absolute endogenous insulin deficiency (168). In addition to these physiological

consequences, hypoglycemia can also have a profound impact on the lives of individuals

with diabetes, and it has been noted that this population may fear hypoglycemia more than

24

the long-term diabetic complications (1;258). As such, the management of glycemia in

diabetes is impeded by the threat of iatrogenic hypoglycemia (161).

Physiological modifications occur following recurrent hypoglycemia. In animal models,

moderate recurrent hypoglycemia has been shown to have a protective role in preventing

age-related decline of hippocampal cognitive function (259) and decreasing brain damage

after severe hypoglycemia (260). However, these beneficial effects do not negate the fact

that compromised counterregulatory hormone defenses against low blood glucose levels

caused by recurrent hypoglycemia persist, which result in psychological and physical

morbidities (162). In diabetes, recurrent hypoglycemia decreases glycemic thresholds for

counterregulatory responses (203) and decreases neuroendocrine and autonomic responses

to subsequent hypoglycemia. In type 1 diabetes, prior, repeated bouts of hypoglycemia

cause further suppression of the already impaired counterregulatory hormone responses

that are observed with the disease (203;205;220;227;261-263). Following recurrent

hypoglycemia, diabetic subjects show markedly diminished epinephrine counterregulation

compared with acutely hypoglycemic controls (203;205;223;226;261;263;264).

Interestingly, this epinephrine defect was not observed in a study that tested recurrent

hypoglycemia with episodes 2 days apart, and the authors suggested that the

pathophysiological effect of recurrent hypoglycemia may be of short duration in patients

with type 1 diabetes (262) as compared to non-diabetic subjects (265). Since glucagon

response is already absent in most diabetic subjects, it has not been shown to be further

worsened by recurrent hypoglycemia (203;264). Similar to that in non-diabetic subjects,

norepinephrine (205;262;264) and glucocorticoid (205;227;264) responses are not as

conclusive since studies have shown either decreased or unchanged counterregulation of

these hormones. Recurrent hypoglycemia in type 2 diabetic individuals with moderate

25

glycemic control has been shown to blunt counterregulation to subsequent hypoglycemia

compared non-diabetic controls, and counterregulatory defenses were further impaired in

intensively treated type 2 diabetes (219). Glucagon and epinephrine hormone

counterregulation, and neurogenic and neuroglycopenic symptom scores to hypoglycemia

were reduced following repeated hypoglycemia in type 2 diabetic subjects, and glycemic

thresholds were also shifted to a lower glycemic level (168).

Ultimately, hypoglycemia remains a serious complication and limiting factor to glycemic

management in both type 1 and type 2 diabetes. Without adequate counterregulation,

recovery of blood glucose levels to euglycemia cannot be achieved, and the result is a

prolonged state of hypoglycemia with the increased potential for morbidity and mortality.

1.7 Somatostatin

In 1968, rat hypothalamic extracts were demonstrated to inhibit growth hormone secretion

from the pituitary gland (266). However, it was not until 1972 that somatostatin, a cyclic

tetradecapeptide that inhibited the release of pituitary growth hormone, was first identified

and sequenced by the laboratory of Guillemin from sheep hypothalami (267). Guillemin was

awarded the Nobel Prize in 1977 for his research on hypothalamic hormones (268). A

second form of the peptide with an extension of 14 additional amino acids at the N-

terminus, somatostatin-28, was subsequently identified (269). The amino-acid sequences

of somatostatin and somatostatin-28 are shown in Figure 1-1. Since its discovery,

somatostatin has been found to be widely distributed throughout the body with a broad

spectrum of biological effects mediated via a family of five receptors. As such, various

analogues for somatostatin designed for therapeutic use began to be synthesized as early

26

as 1974 (270). Several somatostatin-like peptides have also be identified and reviewed and

will be described in brief below.

1.7.1 Functions

The best known physiological role of somatostatin is its regulation of hormone secretion, but

its other diverse effects include regulation of cell growth, nutrient absorption, smooth

muscle contractility, and neuromodulation (271;272). Somatostatin regulates hormone

secretion from the pituitary gland and pancreatic islets, gastrointestinal cell function, and

immune cell function (271).

1.7.1.1 General somatostatin effects in pancreatic islets.

Pancreatic islets have a rich vascular supply and receive up to 10% of the total blood flow to

the pancreas despite the fact that they comprise only about 1% of pancreatic tissue (273).

Somatostatin’s main effect on pancreatic hormones is inhibition of both insulin and glucagon

release (274;275). It also has an autocrine function whereby somatostatin inhibits its own

release (272). Pancreatic -cells are juxtaposed to insulin- and glucagon- containing -cells

and -cells, respectively, within islets (276). There is also evidence of paracrine intraislet

control exerted by somatostatin since -cells are in close proximity to both - and -cells in

rat and human islets and -cell processes were observed to extend into -cell clusters in rat

islets (277;278). Isolated pancreatic islets incubated in a medium containing anti-

somatostatin antibodies to bind endogenously secreted somatostatin have significantly

increased release of insulin and glucagon (279;280), and isolated pancreata perfused with

somatostatin antibody to immnoneutralize islet somatostatin similarly demonstrate

increased stimulation of pancreatic glucagon and insulin release (281;282). Furthermore, in

vivo microscopy experiments of rat and mouse pancreas microcirculation demonstrate a

plausible mechanism in which cell-to-cell communication between -cells and -cells can

27

exist within the islet (283). Taken together, its anatomical juxtaposition to - and -cells,

findings from in vitro and ex vivo somatostatin immunoneutralization studies, and intra-islet

microcirculation experiments all support the concept that somatostatin is a local regulator of

pancreatic islet -and -cell function.

1.7.1.2 Arguments against local somatostatin effects on pancreatic islets.

Some studies suggest that somatostatin-secreting -cells are downstream of glucagon-

secreting -cells and insulin-secreting -cells in the islet microcirculation of non-diabetic

rats, and thus have argued against the regulation of glucagon and insulin by local

somatostatin (79;284;285). This hypothesis of islet blood flow implies that pancreatic

somatostatin must pass through the systemic circulation before acting on cells of the

pancreatic islet. Furthermore, early work with intravenously administered antisomatostatin

serum, which has the effect of neutralizing somatostatin’s actions, demonstrated no

inhibitory effect on plasma insulin and glucagon levels (286), which contrasts the in vitro

and ex vivo effects of antisomatostatin administration and the local role of regulating

pancreatic hormone release. However, it should be noted that antisomatostatin antibodies

are very large in structure, and if given via an intravenous route, may not be accessible to

the interstitium of pancreatic islets for regulation of pancreatic hormone release (88;287).

1.7.1.3 General somatostatin effects in the brain and pituitary gland.

The physiological roles of somatostatin include inhibition of hormone secretion as well as

neurotransmission. Somatostatin inhibits GHRH (288) and TRH (289) secretion from the

hypothalamus. Within the brain, somatostatin can act as a neurotransmitter with effects on

cognition, locomotor, sensory, and autonomic functions (reviewed in: (290-292)). The

localization and release of somatostatin from nerve endings supports the transmitter and

modulator role of somatostatin (293). Notably, somatostatin also modulates the release of

28

other brain neurotransmitters: it inhibits the release of norepinephrine, corticotrophin

releasing hormone (CRH), and endogenous somatostatin from the hypothalamus – all of

which may play a role in the counterregulatory hormone response. Somatostatin inhibits

growth hormone release from the anterior pituitary (267;272). Release of thyrotropic

hormone, also known as thyroid stimulating hormone (TSH), from the anterior pituitary is

inhibited by somatostatin (272). Somatostatin also inhibits the secretion of

adrenocorticotrophic hormone from the anterior pituitary (ACTH) (271;294), but it has not

been shown to affect the reproductive hormones luteinizing hormone (LH), follicle-

stimulating hormone (FSH), or prolactin in normal subjects (271).

1.7.1.4 General somatostatin effects in the gastrointestinal tract.

In the gastrointestinal tract, somatostatin acts as a neurotransmitter in the myenteric

plexus and as a paracrine regulator in epithelial cells where it influences the function of

adjacent cells (272). By and large, somatostatin inhibits gut exocrine secretions of gastric

acid, pepsin, bile, and colonic fluid. Somatostatin also suppresses motor activity of the gut

by inhibiting gastric emptying, gall bladder contraction, and small intestine segmentation

and decreases splanchnic blood flow and glucose absorption from the gut. In general,

somatostatin appears to inhibit the majority of gut hormones, such as vasoactive intestinal

polypeptide, gastrin, gastric inhibitory peptide, secretin and cholecystokinin (271;295).

Somatostatin also inhibits stimulated glucagon-like peptide-1 release from the gut

(296;297).

1.7.1.5 Other effects of somatostatin relating to glucose homeostasis.

Somatostatin has well-established effects on regulating the secretion of hormones

responsible for maintaining glucose homeostasis. But the question arises whether

somatostatin has other direct effects. Somatostatin decreases hepatic glucose output, and

29

this effect is thought to be secondary to the inhibition of pancreatic hormone secretion

(295). In a hyperinsulinemic-euglycemic clamp in non-diabetic humans, somatostatin

infusion at a dose commonly used in human investigation (600 g/h) did not increase

insulin-stimulated glucose uptake in humans (298).

1.7.2 Localization of somatostatin-producing cells

Two main types of somatostatin-producing cells exist and have different morphologies: 1)

secretory cells, which have short cytoplasmic extensions; and 2) nerve cells, with multiple

branches. In the rat, approximately 65% of total body somatostatin-immunoreactivity is

from the gastrointestinal tract, 25% from the brain, 5% in the pancreas, and 5% from other

tissues (271).

1.7.2.1 Pancreatic somatostatin-producing cells.

In the pancreas, somatostatin-producing cells are found in -cells of the islet. Somatostatin

was identified in a population of islet cells that differed from glucagon- and insulin-secreting

cells in 1974 (299). In rodent islets, the proportion of -cells relative to total cells of the

islet (5-10%) is normally far less than that of -cells (60-80%) and -cells (15-20%) but

greater than that of pancreatic polypeptide (PP)-cells and ghrelin-secreting -cells (<3%)

(300-303). The lower proportion of -cells is the same for human islets, in which islet cell

composition differs from rodent islets. Numerous recent reports have indicated that

compared with rodent islets, the percentage of -cells (~50%) and -cells (~40%) is less in

humans, but the percentage of -cells is similar (~10%) (301;304). Some have suggested

that the human islets have as much as twice the proportion of -cells as mouse islets (305).

This proportion of -cells is altered in many models of diabetes, which will be discussed

below. Pancreatic -cells secrete somatostatin into capillaries or perivascular spaces within

islets and normally contribute to approximately 5% of circulating somatostatin (271;306).

30

Approximately 15% of -cells lack cytoplasmic processes, which implicates a paracrine role

for -cells. In the adult, somatostatin-14 is the primary secretory product of pancreatic -

cells (307;308). Individual islet cells bind somatostatin-14 differently from somatostatin-

28. Although somatostatin-14 binds with - and -cells, and somatostatin-28 binds with -,

-, and -cells, there appears to be a preferential binding association of somatostatin-14

with -cells and somatostatin-28 with -cells (309). The differential binding is likely due to

the differential distribution of somatostatin receptors on each islet cell type (310), as will be

discussed below.

1.7.2.2 Brain somatostatin-producing cells.

The brain is rich with somatostatin-positive neurons and fibers (292;311;312).

Somatostatin-cells are abundant in the hypothalamus, brainstem, the deeper layers of the

cerebral cortex, and limbic structures. Hypothalamic somatostatin is most well known for

its role in inhibiting growth hormone and thyroid-stimulating hormone release from the

anterior pituitary (271). Approximately 80% of hypothalamic somatostatin is located in

cells of the periventricular nucleus which project their axons to the median eminence, and

this comprises the main somatostatinergic tract (271;313). Somatostatin-producing

neurons are also located in the paraventricular nucleus (122) which also project to the

median eminence (292); arcuate nucleus; supraoptic nucleus; suprachiasmatic nucleus; and

ventromedial nucleus of the hypothalamus (293;311;312). In the brainstem region,

somatostatin is also localized in the dorsal vagal complex (311;314). Somatostatinergic

neurons of the nucleus of the solitary tract project to the hypothalamic paraventricular

nucleus (292). As discussed in earlier sections, these hypothalamic nuclei and the dorsal

vagal complex are well-known for their role in regulating glucose homeostasis (13;14).

31

Thus, colocalization of somatostatin in these glucose sensing and regulatory brain regions

could implicate a putative role for somatostatin in regulating glucose homeostasis.

1.7.2.3 Gastrointestinal somatostatin-producing cells.

Within the gastrointestinal tract, there are two main types of somatostatin-producing cells:

D-cells in the mucosa of the pyloric antrum and in the intestine and neurons that are part of

submucosa and myenteric plexuses (313;315-317). In the myenteric plexus, somatostatin

inhibits the induced release the neurotransmitter acetylcholine (315).

1.7.2.4 Other somatostatin-producing cells.

In addition to these main organs that secrete most of the body’s somatostatin, there are

other tissue and cell types that also produce somatostatin. Within the thyroid, somatostatin

is produced with calcitonin in a subpopulation of C-cells (313;318). Immune and

inflammatory cells of the spleen and thymus (e.g. lymphocytes, macrophages) also secrete

somatostatin in small amounts when activated (319;320). Somatostatin inhibits the release

of growth factors and cytokines and inhibits the proliferation and adhesion of lymphocytes

(271;321), presumably as a way of regulating the secretions and functions of these cells.

Whereas somatostatin was detected at high levels from extracts of the posterior pituitary,

the anterior pituitary had no detectable levels of somatostatin (313;322). There are also

small numbers of somatostatin-producing cells in the kidneys and in the adrenal glands

(271).

1.7.3 Synthesis and structure

Somatostatin is synthesized from the preprosomatostatin gene (323). The rat somatostatin

gene encodes preprosomatostatin, a precursor of 116 amino acids that is processed within

the endoplasmic reticulum to yield prosomatostatin, a peptide of 92 amino acids (323).

32

Prosomatostatin is subsequently cleaved posttranslationally to produce somatostatin-28 and

somatostatin-14 by prohormone convertases 1 and 2 (PC1/PC2) and carboxypeptidase E

(CPE) (272). The two forms of the peptide are found in varying amounts in somatostatin-

producing cells due to the differential processing by PC1, PC2, and CPE (271).

Somatostatin-14 can also be derived from alternate processing of somatostatin-28.

Somatostatin-14 is the predominant form in the brain, pancreatic islets, retina, peripheral

nerves, and enteric neurons. Somatostatin-28 is predominant form in the gastrointestinal

tract, especially the duodenum and jejunum, where the prosomatostatin gene is processed

in the intestinal muscosal cells. Somatostatin-28 accounts for approximately 20-30% of

total immunoreactive somatostatin in the brain (271).

The preprosomatostatin gene in humans has striking homology with other vertebrates. The

sequence of somatostatin-14 is completely conserved between fish and mammals, but

mammalian somatostatin-28 shares only 40-60% homology with fish somatostatin-28

(324). Two other somatostatin-like peptides have also been identified: cortistatin and

neuronostatin.

1.7.3.1 Cortistatin.

A somatostatin-like gene called cortistatin gives rise to two cleavage products which are

comparable to somatostatin-14 and somatostatin-28: cortistatin-17 (human) or cortistatin-

14 (rat) and cortistatin-29 (human and rat) (325). Cortistatin shares high structural

homology with somatostatin and binds to somatostatin receptor subtypes with similar

affinity, but its actions, tissue expression patterns, and regulation are not completely the

same as that of somatostatin. Cortistatin gene expression was originally thought to be

restricted to the cerebral cortex and hippocampus, but its mRNA expression has since been

detected in peripheral tissues also known to express somatostatin including the pancreas,

33

adrenal, liver, and small and large intestine (326). Its presence in the brain is only at a

small fraction of the brain with far less distribution (272). Cortistatin has been

demonstrated to have some similar hormone regulatory effects as somatostatin (namely,

inhibition of growth hormone and insulin release), but it also has additional distinct

functions on sleep modulation (327), brain function (325), and effects on immune cells

(328).

1.7.3.2 Neuronostatin.

Neuronostatin is a 13-amino acid peptide encoded by the preprosomatostatin gene (329).

Its amino acid sequence shares very little similarity to somatostatin (329). It is highly

produced in the hypothalamus, and neuronostatin-immunoreactive cells were found to be

comparable in number and intensity to somatostatin immunoreactive cells in the

hypothalamic periventricular nucleus. However, these peptides differ in their calcium

mobilizing effects in cultured rat hypothalamic neurons, which results in diverse cellular

activities (somatostatin reduces, while neuronostatin increases, calcium mobilization) (330).

For example, intracerebroventricular treatment with neuronostatin increased blood pressure

but suppressed food intake and water drinking (329). Furthermore, neuronostatin has no

effect on stimulated GH secretion from pituitary. Neuronostatin can also be found in the

pancreas where it is thought to be colocalized with somatostatin in islet -cells.

Importantly, whereas somatostatin activated the Gi-protein signalling pathway mediated via

the five different somatostatin receptors, neuronostatin had no such effect, which likely

indicates that neuronostatin does not bind to native receptors for somatostatin (329).

1.7.3.3 Apelin

Apelin is a circulating and paracrine peptide hormone which also acts as a neuromodulator.

It has various isoforms: apelin-36, apelin-17, and apelin-13 (331). It is secreted by adipose

34

tissue and is also found in the gastric mucosa and hypothalamus. Specifically in the

hypothalamus, apelin is detected in the SON, PVN, and SCN (331). Known for its

hypotensive effects (332) and also as an adipokine (333), apelin has more recently

garnered attention for its pancreatic effects on insulin inhibition (334;335) and for its

localization within the pancreatic islet. Apelin is found in cells and -cells of human, rat,

and mouse islets but not -cells or PP-cells (335). Apelin itself is upregulated directly by

insulin in both human and rodents (336), and islet apelin mRNA expression is suppressed by

glucocorticoids (335). Apelin has been shown to lower plasma glucose via increased glucose

utilization in fat and muscle (337;338). Similar to somatostatin, the receptor for apelin –

the APJ receptor – is a G-protein-coupled receptor which is expressed in various parts of the

brain: hippocampus, striatum, thalamus, cortex, cerebellum, and hypothalamus (332;339).

The APJ receptor is upregulated in the hypothalamic PVN by acute and repeated stress

(340). More recently, APJ expression was found in human and mouse islets (335).

Recently, it was reported that children with type 1 diabetes have increased circulating apelin

levels compared to non-diabetic controls. However, no correlations were found between

elevated apelin levels and body mass index, glycemia, lipids, and insulin sensitivity in these

type 1 diabetic children (341).

1.7.4 Regulation of somatostatin synthesis

The regulatory domains that are located upstream of the 5’ transcriptional unit of the rat

somatostatin gene contain: a TATA box, a cAMP response element (CRE), two nonconsensus

glucocorticoid response elements (GREs), and a consensus insulin response element CGGA

activated by an ETS-related transcription factor (271). Pancreatic duodenal homeobox 1

(PDX-1) is a transcriptional factor required for activating somatostatin gene transcription

and maintains pancreatic islet functions along with activating gene transcription of insulin,

35

islet amyloid polypeptide, glucose transporter type 2, and glucokinase (342). Transcription

of the somatostatin gene can be repressed by two silencer elements located in the promoter

region (343).

Somatostatin mRNA levels are regulated by many of the factors that also influence

somatostatin secretion. Somatostatin gene transcription is stimulated by glucocorticoids,

growth hormone, testosterone, estradiol, NMDA receptor agonists, growth factors (IGF-I,

IGF-II) and cytokines (IL-1, IL-10, TNF-, and IFN-). Conversely, somatostatin gene

transcription is also inhibited by glucocorticoids, insulin, leptin, and TGF-. Glucocorticoids

have a dual effect on the somatostatin gene by enhancing its synthesis via its interaction

with the glucocorticoid receptor and GRE and inhibiting its synthesis via accelerated

somatostatin transcript degradation. Insulin is also known to accelerate somatostatin mRNA

degradation. Among the intracellular mediators known to modulate somatostatin mRNA,

Ca2+, cGMP, nitric oxide, and cAMP, cAMP most potently activates somatostatin gene

transcription. cAMP induces PKA phosphorylation of nuclear cAMP response element binding

protein (CREB), which binds to CRE (271).

In neurons, somatostatin gene transcription is strongly activated by binding of the

phosphorylated transcription factor cAMP response element-binding protein (CREBP) to the

cAMP response element in the promoter sequence of its gene (344;345) and the

homeodomain transcription factor PDX-1 to the promoter element TSEI. In pancreatic

islets, somatostatin gene is also upregulated by cAMP and PDX-1. Interestingly, enhancer

elements in the promoter region of the somatostatin gene that upregulate its expression in

pancreatic islets may actually have the opposite function and act as gene silencer elements

in neurons. This includes complexes that bind transcription factors PAX6, PBX, and PREP1

to TSEII and UE-A elements in the promoter region of the somatostatin gene (272).

36

Somatostatin has a short half-life (approximately 1.1 to 3 minutes in humans) since it is

rapidly metabolized by peptidase enzymes in the blood and tissues. The kidney and liver

are the main sites of somatostatin clearance (271).

1.7.5 Regulation of somatostatin secretion

Some agents exert common secretory effects on somatostatin-producing cells throughout

the body while others have differing effects depending on the type of tissue. Typically, the

divergent effects of these agents occur because of tissue-specific expression of certain

receptors for the agent or because of indirect effects through the release of other

transmitters or peptides (271). Generally, membrane depolarization of somatostatin-

containing nerve cells or peripheral somatostatin secretory cells is a common mechanism

that elicits somatostatin release (291).

1.7.5.1 Regulation of pancreatic somatostatin.

Somatostatin release from pancreatic islets -cells is normally increased in response to

glucose stimulation (305;346) and other nutrients such as amino acids and lipids (271).

Glucose stimulation leads to closure of KATP-channels (presumably via an increase in the

cytoplasmic ATP/ADP ratio), and membrane depolarization. Sulfonylureas also stimulate

somatostatin release by closing KATP channels (347). Glucagon stimulates the release of

somatostatin (348). Insulin inhibits pancreatic somatostatin release (271) as does

activation of gamma-aminobutyric acid (GABA) receptors on -cells (295). Somatostatin

secretion by pancreatic -cells is stimulated by activation of both -adrenergic and

cholinergic receptors (295). In isolated perfused human pancreas, stimulation of the celiac

neural bundle has the predominant effect of inhibiting glucose-stimulated somatostatin

secretion via an -adrenergic mechanism (349). -adrenergic fibers stimulate somatostatin

secretion, whereas cholinergic fibers had a weaker effect (349). Similarly in perfused dog

37

pancreas, somatostatin secretion was increased by -adrenergic activation and suppressed

by -adrenergic activation (72). Interestingly, destruction of the VMH increased pancreatic

somatostatin release to nutrient stimulation (350). CCKB receptors found on pancreatic -

cells (351), and more recently, occupation of CCK-1 and CCK-2 receptors was shown to

initiate somatostatin secretion in rat pancreatic islet cell line RIN-14B cells (352).

1.7.5.2 Regulation of brain somatostatin.

Unlike its action in the pancreas, glucose inhibits hypothalamic somatostatin secretion, and

amino acids do not appear to have a role. Likewise, insulin stimulates hypothalamic

somatostatin, which is opposite to its effect in the pancreatic islet. Somatostatin also has

an autoregulatory effect whereby it inhibits its own release in the hypothalamus (271).

Growth hormone and thyroid hormones enhance somatostatin release, suggesting that

somatostatin plays a negative feedback role on regulating these hormones (353;354).

Secretion of somatostatin from the hypothalamus is promoted by dopamine, substance P,

glucagon, acetylcholine, norepinephrine, vasoactive intestinal polypeptide, and

cholecystokinin (318;355). Various cytokines, such as IL-1, IL-6, and TNF-, also stimulate

somatostatin secretion from the rat brain (356). In contrast, leptin and TGF- have been

shown to inhibit somatostatin secretion from rat hypothalamic cells (357;358). Opiates and

GABA likewise inhibit somatostatin release.

1.7.5.3 Regulation of gastrointestinal somatostatin.

Although the focus of this thesis is not on gastrointestinal somatostatin regulation, it is

important to briefly mention stimulators and inhibitors of somatostatin from the gut since it

represents the main source of circulating somatostatin (271). Gut secretion of somatostatin

is triggered by luminal, but not circulating somatostatin (291;359). Stimulation of the

vagus nerve induced secretion of somatostatin into the antral lumen (360). Somatostatin

38

release is also stimulated by insulin (271) as well as GLP-1, but not GLP-2 (361). Gastrin-

releasing peptide stimulates somatostatin secretion from the intestine, but galanin,

vasoactive intestinal peptide, and epinephrine had no effect (361). Bombesin, a

neuropeptide and hormone that stimulates gastric release and aids in negative feedback

signals to suppress eating, also stimulates somatostatin (362). In the fasting state, plasma

concentration of somatostatin is low (30–100 pg/mL) and increases by approximately 2-fold

in the postprandial state (363). This postprandial increase in plasma somatostatin is mostly

due to somatostatin derived from the intestinal epithelium (364;365).

1.8 Somatostatin receptors (SSTRs)

Somatostatin receptors (SSTRs) were first described in 1978 in pituitary cell lines (366), but

it was not until 20 years after the discovery of somatostatin that structure of one of its

receptors was characterized by molecular cloning (367). It is now known that SSTRs are

found in various densities in the brain, pituitary gland, gastrointestinal tract, endocrine and

exocrine pancreas, adrenal glands, thyroid, kidneys, and immune cells (271). Tumour cell

lines are also rich in SSTRs, but a review of this literature is beyond the scope of the

present thesis. For the purposes of this thesis, the general localization and function of the

family of receptors for somatostatin, SSTR1 to SSTR5, inclusive, will be briefly examined,

and more emphasis will be taken to examine our SSTR of interest, SSTR2.

Somatostatin binds to each of the SSTRs with very high affinity; that is, in the nanomolar

range (368). Somatostatin-14 binds to all the SSTRs with greater affinity with the

exception of SSTR5, in which somatostatin-28 is the ligand with greater selectivity (369).

The receptors can also be grouped into two subfamilies in which SSTR2, 3, and 5 bind well

39

to short somatostatin analogues (i.e. hexapeptide and octapeptides) and SSTR1 and 4

which have poor binding with these short peptide analogues (369). In general, the SSTR

subtypes are of a similar size (356-391 amino acids in length), and each has seven -helical

transmembrane segments typical of G-protein coupled receptors with an extracellular N-

terminus and an intracellular C-terminus. In the seventh transmembrane segment, there is

a signature sequence for this family of receptors that is highly conserved in all SSTR

isoforms from humans and other species. The five SSTR subtypes display a high level of

structural conservation across species (369;370). The nearest related family of receptors to

SSTRs are the opiod receptors (371).

1.8.1 General localization of SSTRs

The central nervous system, pituitary, and gastrointestinal tract ubiquitously express all

members of the SSTR family, SSTR1-5. In the central nervous system, SSTR1 is most

highly expressed in the neuraxis (the axis of the central nervous system) and throughout

the entire brain. SSTR2 has a high level of expression in the cerebral cortex. SSTR3 has

preferential expression in the cerebellum, SSTR4 is less well expressed as compared to the

other types and in early studies was referred to as the “brain-specific” SSTR (372), and

SSTR5 is moderately well expressed throughout the brain (373-377). Interestingly, brain

SSTR5 in humans is less well expressed than in rats (376). Specifically in the

hypothalamus, SSTR1 has the highest overall expression, followed by SSTR2, 4, 3 and 5.

SSTR1 seems to be highest expressed in neurons of all major hypothalamic nuclei, in nerve

fibers along the median eminence, and along the third ventricle. SSTR2 is likewise

expressed in these areas but at a lower density than SSTR1. SSTR3 is localized in the

paraventricular nucleus, dorsomedial nucleus, arcuate nucleus, and median eminence.

SSTR4 is localized to the arcuate nucleus, ventromedial nucleus, and median eminence.

40

SSTR5 is the least expressed subtype and appears in the median eminence (374;375;378).

In the pituitary, all five subtypes of SSTR are expressed in rats, but human pituitaries lack

SSTR4. SSTR5 is the most predominant subtype expressed in the pituitary, followed by

SSTR2 (373;379-382). In the gastrointestinal tract, all five SSTRs have been identified in

the stomach and small and large intestine, and the enteric plexus expresses SSTR1-3

(383;384).

Rat and human pancreatic islets also express all five SSTR subtypes, but the species differ

in their level of SSTR expression on the different islet cell types. In human pancreatic islets,

insulin-secreting -cells highly express SSTR1 and SSTR5, glucagon-secreting -cells

predominantly express SSTR2, and somatostatin-secreting -cells predominantly express

SSTR5 (385). In human isolated islets, it was demonstrated that SSTR2 was the

predominant subtype that mediates the suppression of glucagon release by somatostatin

(386). In rodent islets, the main difference is that -cells more exclusively express SSTR5

(387). Rat -cells and -cells show the same selective predominant expression of SSTR2

and SSTR5, respectively (388-391). Functional studies in rodents are consistent with the

above findings of receptor localization showing that SSTR2 mediates the inhibitory effect of

somatostatin on glucagon secretion while SSTR5 inhibits insulin release (388;389;392-394).

Notably, one group has reported slightly different findings in which rat and mouse -cells

showed high expression of SSTR2 and SSTR5 and - and -cells showed high levels of

SSTR1, 2 and 5, suggesting that more than one SSTR needs to be activated to suppress

islet glucagon and insulin release (395;396).

In the adrenal gland, there is a high concentration of SSTR2 (390;397-399) and also

moderate levels of SSTR1 and SSTR3 (373;400). In the liver and kidney, SSTR3 is

expressed. The lung, heart, and placenta express SSTR4. Immune cells preferentially

41

express SSTR2 (319;401;402), but recent studies report that SSTR1 and SSTR2 colocalized

on human macrophages are responsible for the immunosuppressive effect of somatostatin

(403). The retina expresses all SSTRs, with highest levels of SSTR1 and SSTR2 (404).

Interestingly, the majority of human tumour cells are usually positive for SSTRs (271) with

predominant highest frequencies reported for SSTR2 (405) or SSTR3 mRNA expression

(406-408). More specifically in human neuroendocrine tumours, a recent study

demonstrated that SSTR1 and SSTR5 were the most highly detected forms of receptors,

followed by SSTR3 and SSTR2 (409).

1.8.2 General functions of SSTRs

SSTRs mediate the functions of somatostatin. As previously mentioned, one of the main

functions of somatostatin is to inhibit exocytosis in target cells. This action is mediated

predominantly via SSTR2, SSTR3, and SSTR5 and involves i) decreased intracellular cAMP

due to inhibition of adenylyl cyclase, ii) decreased Ca2+ influx due to action on K+ and Ca2+

ion channels, and iii) stimulation of phosphatases such as calcineurin (which inhibits

exocytosis) and serine threonine phosphatases (which dephosphorylate and activate K+ and

inhibit Ca2+ ion channels) (369). In addition to its anti-secretory effects, somatostatin also

has: 1) anti-proliferative, 2) anti-growth, and 3) apoptotic functions (271). The anti-

proliferative effects of somatostatin are mediated primarily via SSTR1, 2, 4, and 5 by which

it induces cell cycle arrest at the G1 phase. Anti-proliferative effects of somatostatin have

been demonstrated in normally dividing cell types such as activated lymphocytes and

inflammatory cells (410;411), intestinal muscosal cells (318), bone and cartilage cells

(412), as well as tumour cells (271). SSTRs activate phosphotyrosine phosphatases (PTPs)

to transduce anti-growth signals via modulation of a mitogen-activated protein kinase

(MAPK) pathway. SHP-1 and SHP-2 belong to the family of PTPs which are implicated in

42

SSTR-mediated anti-growth signaling (413). Somatostatin induces growth arrest by

indirectly activating nitric oxide synthase via SSTR2 and via a negative coupling with

guanylate cyclase via SSTR5. Somatostatin’s anti-growth effects are mediated via SSTR2

(in various tumour cells) and SSTR4 (in human B lymphocytes) through a mechanism

involving MAPK-ERK1/2 (extracellular regulated kinase) upregulation of cyclin-dependent

kinase inhibitors (414). Via SSTR3, somatostatin activates apoptosis by inducing p53 and

Bax expression (415).

1.8.3 Specific localization, ligand binding, and signalling of SSTR2

The rat SSTR2 gene was identified in 1992 (416). Two spliced variants of the SSTR2 gene

exist: a long form (SSTR2A), and a short form (SSTR2B). These two variants are generated

through alternate mRNA splicing and differ only in the length of their cytoplasmic tail, with

the SSTR2B form differing in 15 amino acids at the C-terminus (400;417). The C-terminal

tail is not essential for coupling to G-proteins or adenylyl cyclase. There is a 93-96%

conservation of the sequence identity of SSTR2 isoforms among human, rat, mouse,

porcine, and bovine species.

SSTR2 is highly prevalent in the kidney, and pancreas and along with SSTR1 is prevalent in

the stomach, pituitary, small intestine (particularly the duodenum), and spleen

(271;367;385). It is the main subtype expressed in lymphocytes and inflammatory cells

(319;401;402). There are also low levels of SSTR2 detected in the liver (367) although in

healthy liver, SSTR2 may be localized on other cell types such as cholangiocytes rather than

hepatocytes (418). SSTR2 is the main subtype responsible for the suppression of gastric

acid (388).

43

In human pancreatic islets, SSTR2 is predominantly expressed on glucagon-secreting -cells

(385). In human isolated islets, it was demonstrated that SSTR2 was the predominant

subtype that mediates the suppression of glucagon release by somatostatin (386). Rat -

cells show the same selective predominant expression of SSTR2 (388-391). Functional

studies in rodents are consistent with the above findings of receptor localization showing

that SSTR2 mediates the inhibitory effect of somatostatin on glucagon secretion

(388;389;392-394).

In the brain, moderate to high levels of SSTR2 expression include the basolateral amygdala

(BLA) (419-421); in neurons of all major hypothalamic nuclei (378;422), including

mediobasal hypothalamus (312;423) and specific nuclei regulating endocrine and fluid

homeostasis such as the supraoptic nucleus (SON) and PVN and feeding centers such as the

ARC and dorsomedial nucleus (420;424-428); cerebral cortex (419;420); parabrachial

nucleus, locus coeruleus, and NTS of the brainstem (419;420), and in the nerve fibers along

the median eminence (378). Intracerebroventricular administration of a selective SSTR2

agonist elicited neuronal activation (as measured by an increase in Fos expression) in brain

regions such as the BLA, SON, PVN, ARC, parasubthalamic nucleus, lateral parabrachial

nucleus (429). A recent study reports that selective activation of brain SSTR2 induces a

feeding response while blocking brain SSTR2 reduced food intake (430). Interestingly, the

effect of this antagonist was specific to central administration and not reproducible by

intraperitoneal administration. Drinking, grooming, and locomotor activity were also

increased by selective activation of brain SSTR2 (430).

With SSTR5, SSTR2 inhibits growth hormone and thyroid stimulating hormone from human

and rat pituitary cells (431;432). However, there is also evidence demonstrating the

importance of SSTR1 in regulating GHRH and growth hormone release (423;428;433;434).

44

SSTR2 is not internalized as readily as the other SSTRs (435-437). From studies that

demonstrated that SSTR2 is sensitive to GTP analogues and pertussis toxin, it is now known

that SSTR2 is associated with inhibitory G-proteins Gi and Go, which are the G-proteins that

involved with the inhibition of adenylyl cyclase (438).

1.9 Somatostatin in type 1 diabetic humans and animal models

Somatostatin levels are increased in many diabetic subjects studied. The most notable

change is the increase in pancreatic somatostatin content which is has been observed in

type 1 diabetic humans (439) in which an increased number and volume of -cells has been

observed (440). Pancreatic somatostatin release was elevated in type 1 diabetic subjects

(363;441), particularly in those with poor glycemic control (441). Increased basal plasma

somatostatin has also been reported in type 1 diabetic individuals (442;443). Experimental

diabetes induced in monkeys (444) showed increased -cell volume and hyperplasia.

Alloxan-diabetic dogs also demonstrated elevations in islet somatostatin content and -cell

area (90;445) as well as increased basal and nutrient-induced circulating somatostatin

levels (446) as compared to non-diabetic dogs.

There is an abundance of literature demonstrating increased somatostatin levels in

streptozotocin- (STZ-) diabetic rats. STZ-diabetes is an experimental model of diabetes

(discussed in section 3.1.2). STZ-diabetic rats have increased pancreatic somatostatin

(174;439;447-452) and secrete more pancreatic somatostatin than non-diabetic controls

(451). It was demonstrated that pancreata from STZ-diabetic rats had more pancreatic

somatostatin content which increased in proportion to the dose of STZ used to induce

diabetes while graded reductions of insulin content were observed (453). In addition, these

45

pancreata had increased arginine-stimulated secretion of somatostatin, which also

negatively correlated with insulin release. This suggests that pancreatic somatostatin

content and secretion are proportional to the state of insulin deficiency (453). Pancreatic

prosomatostatin mRNA was increased in STZ-diabetic rats compared to non-diabetic rats

(205;454). Plasma somatostatin levels were also elevated in STZ-diabetic rats

(174;205;455). Fasting reduced basal plasma somatostatin in both STZ-diabetic and non-

diabetic rats (455). The stomach of STZ-diabetic rats was also shown to have elevated

levels of somatostatin as compared to non-diabetic rats (452;456;457). These D-cells of

STZ-diabetic rats also secreted more somatostatin when stimulated than those of non-

diabetic counterparts (456). Small intestine somatostatin content was also increased in

STZ-diabetic rats compared with control animals (457). Intravenous arginine increased

plasma somatostatin levels to a greater extent in STZ-diabetic rats as compared to non-

diabetic rats (455).

Alloxan-diabetes is another form of induced, experimental diabetes. In alloxan-diabetic

rats, pancreatic -cell mass was increased (458;459), as was arginine-stimulated

somatostatin release (459). In one study, hyperplasia of -cells was observed after 14

months of alloxan diabetes (458).

Increased pancreatic somatostatin has also been reported in NOD mice (460). In NOD mice

15 days after onset of diabetes, pancreatic islets were characterized by infiltration of

lymphocytes, and somatostatin-containing cells occupied the majority of endocrine cells of

the islets while insulin-containing cells decreased as expected and glucagon-containing cells

remained similar (460).

46

There is evidence in animal models of diabetes that insulin treatment can reduce and/or

partially normalize pancreatic somatostatin levels (90;351;452), arginine-induced elevations

of plasma somatostatin (455), and high basal somatostatin secretion rates in islets from

untreated diabetic rats (461). This suggests that that increased somatostatin during

diabetes may be due, at least in part, to a lack of endogenous insulin action or its metabolic

consequences. However, it should be noted that insulin treatment of type 1 diabetic

humans did not prevent the abundance of -cells found in human diabetic pancreata (439)

nor the elevation in plasma immunoreactive somatostatin levels (363). In addition, mRNA

expression of pancreatic prohormone convertases PC1 and PC2 were found to be >2-fold

and >4-fold increased, respectively, in STZ-diabetic rats, which may also be in part

responsible for increased pancreatic somatostatin in diabetes (462). It has also been

postulated that increased glucagon, representing an absolute or relative glucagon excess,

within the pancreatic islet might lead to a compensatory response of the -cell to secrete

more somatostatin. This postulate was supported by observations that glucagon-secreting

-cell tumours also showed hyperplasia of -cells (439).

Whereas glucose stimulates somatostatin-secretion from healthy -cells, somatostatin-

secreting -cells from islets from STZ-diabetic rats were unresponsive to changes in glucose

concentrations but still responded to increased cAMP levels (461). This suggests that the

hyperglycemia associated with diabetes may not be responsible for the increased pancreatic

somatostatin release observed in diabetes. In vivo insulin treatment of STZ-diabetic rats

restored the glucose sensitivity of -cells (461). Literature also widely supports the concept

that somatostatin release is stimulated by glucose and tolbutamide in healthy individuals,

but that these effects are lost in type 1 and type 2 diabetes, which has been proposed to

47

contribute to the impaired prandial suppression of glucagon secretion in diabetes (305;463-

468).

1.9.1 Brain somatostatin in experimental diabetes

The literature regarding brain somatostatin levels is less consistent than that of the

pancreas and gastrointestinal tract. In untreated STZ-diabetic rats, somatostatin mRNA

expression was reduced in the central portion of the periventricular nucleus with no changes

observed in the other somatostatin-rich hypothalamic areas: the periventricular nucleus or

the arcuate, suprachiasmatic or paraventricular nuclei (469). Interestingly, this anatomical

area of somatostatin reduction is involved with regulation of the anterior pituitary hormone

release. These changes in hypothalamic somatostatin expression were restored with insulin

treatment that normalized blood glucose levels (469). Older studies, however, did not

observe changes in hypothalamic somatostatin content or mRNA expression in STZ-diabetic

as compared to non-diabetic rats (448;449;452;470). Similarly, there was no change in

hypothalamic somatostatin content of db/db and ob/ob mice (471). It is possible that no

changes were observed if total hypothalamic content were measured as opposed to specific

anatomical areas. Interestingly, others have observed increased hypothalamic secretion of

somatostatin into the hypophyseal portal blood in vivo in STZ-diabetic rats (472). In other

areas of the brain, decreased somatostatin mRNA expression in the hippocampus and

frontal cortex of STZ-diabetic rats has also been reported (473), but how and whether these

anatomical changes affect glucose homeostasis in diabetes has yet to be determined.

48

1.10 Somatostatin in type 2 diabetic humans and animal models

Reports of somatostatin levels and -cell function in type 2 diabetes are not as consistent as

that described in type 1 diabetes. Pancreatic somatostatin secretion was shown to be

greater in type 2 diabetic humans, to a similar extent as type 1 diabetic subjects, as

compared to non-diabetic individuals (441). The number of somatostatin-expressing -cell

was increased by 58% in type 2 diabetic human pancreases as compared with non-diabetic

human pancreases and represented an absolute increase in number (474). However, other

studies showed no difference in percentage volume (475) or number (440) of -cells in type

2 diabetic human pancreatic islets or in basal plasma somatostatin levels (476).

In db/db mice, increased pancreatic somatostatin content and -cell hyperplasia were

observed. In ob/ob mice, pancreatic somatostatin content was also increased, but no

increase was noted in the number -cells (471). However, other studies in db/db and ob/ob

mice demonstrated that there were no differences in hypothalamic, gastric, or pancreatic

somatostatin levels in these genetically diabetic and/or obese mice until their serum insulin

levels decreased (477). Based on the temporal relationships of pancreatic somatostatin to

the disturbances in carbohydrate metabolism, the authors suggest the concept that changes

in pancreatic somatostatin represent a response to, rather than a cause of, hypoinsulinemia

(477).

1.11 Diabetes-induced changes in SSTR expression

Some of the studies that investigated changes in SSTR levels in diabetic animal models had

the purpose of examining growth hormone regulation and thus focused their investigations

on the brain and pituitary gland. Some older studies did not discern between which subtype

49

of SSTR was detected. In STZ-diabetes of short duration (5 and 9 days of diabetes),

concentrations of SSTRs in the hypothalamus and anterior pituitary were greatly reduced

and were not corrected by insulin treatment (478). More specifically, one group showed

that hypothalamic mRNA expression of SSTR5, but not SSTR1-4, and pituitary mRNA

expression of SSTR1, 2, 3, and 5, but not SSTR4, was reduced in STZ-diabetic rats (479).

In these animals, insulin treatment restored hypothalamic SSTR5 levels and partially

restored pituitary SSTR1 and SSTR5 levels. It was suggested that chronic exposure to high

somatostatin levels in diabetes may account for reduced SSTR mRNA expression due to

receptor desensitization, and insulin, high blood glucose levels, or glucocorticoids may also

in part regulate the expression of hypothalamic and pituitary SSTR mRNA expression (479).

However, the explanation behind the subtype-specific reduction is not yet apparent. SSTRs

in the cerebral cortex were not affected by STZ-diabetes of short duration (478).

1.12 Somatostatin: hypoglycemia-induced changes

Circulating somatostatin levels increase in response to insulin-induced hypoglycemia. This

has been observed both in non-diabetic and type 1 diabetic patients without diabetic

autonomic neuropathy (236;480). However, plasma somatostatin levels are not elevated in

response to hypoglycemia in subjects with diabetic autonomic neuropathy, thus

underscoring the importance of autonomic nervous system activity in pancreatic hormone

regulation during hypoglycemia (480). The increase in circulating somatostatin to insulin

hypoglycemia is mediated via activation of the vagus nerve and is in part regulated by the

associated increase in gastric acid (481;482). Truncal vagotomy also abolished the rise in

somatostatin to insulin-induced hypoglycemia in humans. It was concluded that the

somatostatin response is dependent upon vagal integrity and that section of the vagus

50

nerve unmasks a suppressive effect of insulin action or its metabolic or hormonal

consequences on circulating somatostatin levels (483). There is also evidence that

activation of the -adrenergic system by hypoglycemia may stimulate increased secretion of

somatostatin (451;484). Under low glucose conditions, locally secreted glucagon was

shown to increase somatostatin secretion in isolated islets via L-glutamate, a co-secretory

product from -cells that acts on a glutamate receptor on -cells, at least in islets on non-

diabetic rats (485). This response of somatostatin to hypoglycemia is not dependent on

residual -cell function since this rise in somatostatin was demonstrated in both type 1

diabetic patients with and with residual -cell function (486). This rise in plasma

somatostatin was similar in magnitude in both non-diabetic and diabetic individuals (486).

Although insulin-induced hypoglycemia elicited an increase in plasma somatostatin, there

was no rise of somatostatin to surgical stress, thus it has been suggested that this elevation

of somatostatin is likely specific to metabolic stress in non-diabetic humans (487).

Circulating somatostatin levels increase with insulin-induced hypoglycemia but not with

insulin infusion with dextrose to maintain euglycemia, thus this rise in plasma somatostatin

is not due to the effect of insulin but is stimulated indirectly via hypoglycemia (488). In

type 2 diabetic individuals, no rise in plasma somatostatin was observed in response to

insulin-induced hypoglycemia although an increase was observed in non-diabetic individuals

with equivalent hypoglycemia (476).

It is important to note that one of the problems in interpreting plasma somatostatin

responses is the origin of the circulating somatostatin since it can originate from various

tissues (318;489), especially since stimuli such as insulin, glucose, and hypoglycemia can

have divergent effects on hypothalamic and pancreatic somatostatin release, as mentioned

in an earlier section.

51

1.12.1 Hypoglycemia-induced somatostatin and SSTR changes in the brain

Interestingly, hypothalamic somatostatin mRNA expression levels were reported to be

dramatically increased by insulin-induced hypoglycemia in vivo (490). In numerous in vitro

studies, both low glucose concentrations and intracellular glycopenia induced with 2-deoxy-

D-glucose markedly stimulated somatostatin release from rat hypothalamic fragments (491-

494). Glucopenia induced by 2-deoxy-D-glucose, however, did not alter hypothalamic

somatostatin gene expression (494).

In contrast, it has been demonstrated that severe hypoglycemia of a longer duration than

the studies described above produces a disappearance of somatostatin-like

immunoreactivity in the dorsal hippocampus and frontal cortex (495). Furthermore, other

earlier studies demonstrated that hypoglycemia did not affect somatostatin levels in the

hypothalamus, cerebral cortex, hippocampus, and striatum (496-498). However, the total

number of SSTRs (subtype not specified) decreased in the hippocampus and striatum in

response to insulin-induced hypoglycemia as compared to controls but was unchanged in

the hypothalamus and cerebral cortex (496;497).

1.13 Somatostatin and its analogues in diabetic therapy

Type 1 and type 2 diabetic individuals have a decreased secretory response of somatostatin

to glucose stimulation, which has been postulated to contribute to impaired prandial

glucagon inhibition (305;463-468), and hyperglucagonemia can stimulate hepatic glucose

output, thus worsening diabetic hyperglycemia (463;464;467). Thus, somatostatin has also

been considered as an adjunct therapy with insulin for the treatment of diabetes

(489;499;500). The mechanism for improved glycemic control with somatostatin was

52

based on the somatostatin’s action to suppress glucagon and growth hormone secretion and

delay carbohydrate absorption (489). Due to the short half-life of the somatostatin peptide,

synthetic analogs, such as octreotide, were synthesized (501-503). However, somatostatin

analogues have not consistently been shown to be a beneficial adjunct therapy for diabetes

(504;505).

1.14 Somatostatin antagonist – PRL-2903

Functional biochemistry studies have shown that four amino acids (Phe-Trp-Lys-Thr) in the

native somatostatin peptide form a -turn that is essential for the biological activity of

somatostatin (506;507). With this knowledge, various agonists and antagonists with

greater metabolic stability and/or greater binding affinities have been synthesized with

pharmacological selectivity for particular SSTRs.

The SSTR2 peptide antagonist used in the present thesis is PRL-2903 and has the chemical

structure H-Fpa-cyclo[DCys-Pal-DTrp-Lys-Tle-Cys]-Nal-NH2 (Figure 1-1, Figure 1-2). This

peptide was synthesized by Dr. David Coy at Tulane University, Head of Peptide Research

Labs (New Orleans, LA, USA) (508;509). It is also referred to as DC-41-33 and BIM-23458.

This octapeptide has a molecular weight of 1160 da. The peptide has an IC50 of 2.5 nM in a

rat pituitary growth hormone in vitro antagonist assay versus somatostatin (1 nM) and it

binds to cloned human SSTR2 with a Ki of 26 nM. This peptide was also selective for SSTR2

over SSTR3 and SSTR5 by ~10- and 20-fold, respectively, and had negligible binding

affinity to SSTR1 and SSTR4 (509).

53

Figure 1-1. Linear structures of somatostatin, somatostatin-28, and SSTR2 antagonist PRL-2903.

H2N

NH

HN

NH

HN

NH

NH

O

O

O

O O

HN

NH2

O

O

NH2

NF

S S

HN

O

Figure 1-2. Structure of SSTR2 antagonist PRL-2903.

54

54

Chapter 2 Objectives

2 General Objectives

Hypoglycemia can have a profound impact on the lives of individuals with diabetes, and it

this population may fear hypoglycemia more than the long-term diabetic complications

(1;258). Thus, minimizing the incidence of hypoglycemia will improve the quality of life of

people with diabetes until a cure or prevention of diabetes is found.

The main objective of this thesis was to determine a means to improve the

counterregulatory response to hypoglycemia – in particular the glucagon response, which

plays a key role in promoting glycemic recovery (510), and which has consistently been

reported to be impaired in diabetes (167;168;511-513).

2.1 Specific Aim: Study 1

As described in the Introduction and will be further discussed in the chapters ahead, it is

recognized that the impairment in the glucagon response to hypoglycemia in diabetes can

be due to several factors. In brief, these mechanisms are related to local effects in the

pancreatic islet (76), or to central effects in brain regions known to detect glucose levels

and activate glucoregulatory responses (13;193). In addition to these reported factors, we

hypothesized that in diabetes during hypoglycemia, glucagon is suppressed by the

prevailing inhibitory effect of excessive somatostatin acting via SSTR2. Somatostatin has

55

inhibitory effects on the stimulated release of glucagon in the pancreas via SSTR2.

Furthermore, somatostatin and SSTR2 are found in brain regions that regulate glucose-

sensing and glucoregulatory response, and somatostatin is also known to have inhibitory

actions on the secretion of neuropeptides that are involved with the activation of

counterregulatory hormone responses to hypoglycemia. Thus, the first objective of this

thesis was to determine whether defective glucagon, and other hormone,

counterregulation can be restored in diabetic rats if somatostatin actions are

inhibited by using a selective SSTR2 antagonist.

2.2 Specific Aim: Study 2

In light of the fact that we were able to demonstrate a restoration of the glucagon response,

and markedly improved corticosterone response, to hypoglycemia in diabetic rats, we then

endeavoured to determine whether such hormone improvements could translate to

enhanced recovery of hypoglycemia, or prevention of hypoglycemia. From our first study,

we demonstrated deficient glucagon and corticosterone counterregulation to hypoglycemia

in diabetic rats. However, in this model of diabetes, the epinephrine response was not

impaired. Thus, we wanted to test our hypothesis of counterregulatory improvement in a

model with a well-known epinephrine counterregulation defect in addition to impairments of

glucagon and corticosterone – recurrent hypoglycemia (203;205;223;226;261;263;264).

We hypothesized that blocking the actions of somatostatin via SSTR2 would promote

glycemic recovery. Thus, the second objective of this thesis was to determine whether

counterregulatory responses can also be restored by SSTR2 antagonist in diabetic

rats exposed to recurrent hypoglycemia and if such improvements can lessen the

severity of subsequent hypoglycemia.

56

56

Chapter 3 General Materials and Methods

3 General Materials and Methods

3.1 Experimental animals

3.1.1 Care and maintenance

Male Sprague-Dawley rats (Charles River Laboratories, Saint-Constant, QC, Canada) with an

initial body weight of 275-300 g were used. Rats were individually housed in opaque cages

in temperature (22-23°C) and light-controlled (12-h light:12-h dark cycle; lights on

between 0700 and 1900) rooms. Rats had free access to drinking water and fed ad libitum

with rat chow (Teklad Global 18% protein, Harlan Laboratories, Madison, WI, USA). The

rats were allowed 1 week to acclimatize to experimenter handling and their environment

before experimental manipulation. Blood glucose, body weight, and food intake were

monitored for a minimum of 5 days per week for the duration of the experiment. Morning

(fed state) blood glucose was measured using a handheld glucometer (Ascencia Elite

handheld glucometer, Bayer Canada Ltd., Etobicoke, ON, Canada) via a tail-nick blood

sample. All procedures were in accordance with Canadian Council on Animal Care

Standards and were approved by the Animal Care Committee of the University of Toronto.

3.1.2 Streptozotocin-diabetic rat model of diabetes mellitus

Streptozotocin (STZ) consists of a 2-deoxy-D-glucose molecule substituted at C2 with a N-

methyl-N-nitrosourea group (514). The structure of STZ is shown in Figure 3-1 (514). STZ

is a diabetogenic agent used to induce experimental diabetes in animals via its cytotoxic

57

effects on pancreatic -cells (515). The deoxyglucose moiety of STZ directs it to the

pancreatic -cell where it is taken up via glucose transporter GLUT2 (516). The expression

of GLUT2 on -cells is needed for the diabetogenic action of STZ (517). The nitrosurea

moiety of STZ confers its cytotoxic effect.

It is suggested that -cell death is the result of one or more of the following mechanisms: 1)

DNA alkylation; 2) nitric oxide (NO formation); 3) free radical formation. It is proposed that

the primary mechanism for STZ-induced -cell death is DNA fragmentation resulting from

DNA alkylation, which related to the nitrosurea moiety of STZ (518). DNA is alkylated by

highly reactive carbonium ions (CH3+) are formed by the decomposition of nitrosurea (519).

NO is liberated when STZ is metabolized and is known to have cytotoxic effects on -cells

(520). NO also partially mediates a restriction of mitochondrial ATP generation (516). The

reduction of ATP generation results in high intracellular levels of ADP which causes

mitochondrial damage and the generation of hypoxanthine. Xanthine oxidase (XOD), the

activity of which is high in -cells, catalyzes the oxidation of hypoxanthine to xanthine and

xanthine to uric acid, both steps yield hydrogen peroxide and free oxygen radicals as

byproducts. Since -cells are deficient in superoxide dismutase, they are rendered

inefficient at scavenging free radicals and are thus more susceptible to the effects of STZ

(519). Along with DNA alkylation, the production of NO and reactive oxygen species further

contribute to DNA damage, which ultimately leads to an excision and repair process by poly

Figure 3-1. Structure of streptozotocin; see text for reference.

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(ADP-ribose) synthase. This leads to the depletion of cellular nicotinamide adenine

dinucleotide (NAD+), a substrate for poly (ADP-ribosylation) and further reduction of cellular

ATP, which results in the subsequent inhibition of insulin synthesis and secretion (521).

To summarize, the toxicity of STZ is especially relevant to -cells due to several intrinsic

properties of -cells: 1) intrinsically low levels of protective superoxide dismutase; 2)

intrinsically low content of the substrate NAD+, which is needed for DNA repair; 3)

intrinsically high activity of XOD, which forms free radicals; and 4) intrinsic expression of

GLUT2, which is the transporter by which STZ enters the -cell. Thus, -cells are extremely

vulnerable to STZ-toxicity. STZ-induced experimental diabetes is sometimes criticized for

its non--cell effects. Some have argued that other GLUT2-expressing tissues, such as

hypothalamic astrocytes (522), may be compromised by STZ. Nevertheless, the

administration of STZ in rats generates metabolic and endocrine changes that are relevant

for the present study and remains a good a model for diabetes. Approximately 70% of -

cells are destroyed with a single, moderate dose of STZ (65 mg/kg) (470), and thus in

contrast to fully developed type 1 diabetes in which endogenous insulin secretion is

completely absent (523), this form of diabetes is less severe. STZ at 65 mg/kg yields a

diabetic model characterized by fasting and fed hyperglycemia, normal fasting plasma

insulin levels, but reduced fed-state plasma insulin levels (524).

3.1.3 Induction of diabetes using streptozotocin

Streptozotocin (STZ, 65 mg/kg, Sigma Chemical Co., St. Louis, MO, USA) was dissolved in

0.9% saline and immediately injected intraperitoneally to induce diabetes in the rats.

Within 24 h of STZ injection, post-prandial plasma glucose levels were elevated, food intake

was increased, and body weight was less than that of saline-injected controls. Rats that did

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not exhibit hyperglycemia (i.e. > 10 mM) 48 h following injection of STZ were excluded

from the study.

3.1.4 Chronic vessel catheterization

Chronic, indwelling catheters were implanted in the left carotid artery and right jugular bein

to allow for undisturbed blood sampling and re-infusion of red blood cells. Surgery was

performed in designated operating rooms at the University of Toronto’s Department of

Comparative Medicine animal facility under aseptic conditions. All procedures were in

accordance with Canadian Council on Animal Care Standards and were approved by the

Animal Care Committee of the University of Toronto.

3.1.4.1 Pre-operative preparation.

Surgical instruments, gauze, cloths, and gloves were autoclaved prior to use. Sterile, new

syringes, needles, and sutures were used. Catheters were made from polyethylene tubing

(PE-50, length: 80 cm, ID: 0.58 mm, OD: 0.965 mm, Becton Dickinson, Sparks, MD, USA)

with a 3.0 cm- overlap of silastic tubing (ID: 0.76 mm, OD: 1.65, Dow Corning Corporation,

Midland, MI, USA). The silastic overlap was dipped in diethylether to expand the tubing

prior to being slipped over the end of the polyethylene catheter, forming an overlap of 1.5

cm. The silastic end of the catheter was then bevelled at 45° to facilitate insertion into the

vessel. Catheters were soaked in 70% ethanol to disinfect for at least 1 hour, flushed with

saline, and primed with 10 USP/mL heparinized saline (Heparin LEO; 1000 USP units/mL;

LEO Pharma Inc., Thornhill, ON, Canada) prior to vascular insertion.

3.1.4.2 Surgical procedure.

Rats were anesthetized using a ketamine cocktail, which consisted of ketamine (100 mg/kg,

ip; MTC Pharmaceuticals, Cambridge, ON, Canada), acepromazine (1 mg/kg, ip; Wyeth-

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Ayerst Canada Inc., Montreal, QC, Canada), and xylazine (1 mg/kg, ip; Bayer Inc.,

Etobicoke, ON, Canada). Once rats were anesthetized, the ventral side of the neck and the

dorsal side of the neck between the scapulae were shaved and cleaned with 70% ethanol

and 10% povidone-iodine (Betadine solution, Purdue Pharma, Pickering, ON, Canada). The

rat was placed in the dorsal recumbent position on an operating tray, and its chin and limbs

were fastened to the tray with masking tape. The rat was covered with sterile surgical

cloths exposing only the incision area. The ventral skin just anterior to the collarbone was

cut with a 2-cm transverse incision. Using blunt dissection through muscle layers, the left

carotid artery was isolated from the vagus nerve and connective tissue. The exposed vessel

was ligated at the cranial end using sterile 3-0 silk thread. Another ligature was loosely

tied at the caudal end of the exposed carotid. To temporarily stop blood flow to this

segment of the vessel, the exposed carotid artery was pulled taut at the 2 ligatures.

Scissors were used to make a small incision in the vessel wall. The primed, bevelled silastic

end of the catheter was inserted through the opening in the carotid, past the loose caudal-

end ligature, until the silastic-polyethylene overlap of the catheter was also inside the

vessel. The catheter was then secured by tightening the loose ligature at the caudal end.

Two additional sutures were tied to anchor the catheter to the vessel. The catheter was

then tested to ensure blood withdrawal and infusion were possible and that air emboli were

absent. Cannulation of the right jugular vein was carried out using the same procedure.

A 2-cm transverse incision was made through the shaved dorsal surface between the

scapulae. A small subcutaneous pocket anterior to the incision was formed by using forceps

to loosen the skin from the underlying connection tissue. Using a 16G needle and scalpel, a

small opening was made in the skin 1.5 cm anterior to the dorsal incision. A metal coil

tether (rodent tether, 18”; Lomir Biomedical Inc., Notre-Dame-de-I'Île Perrot, QC, Canada)

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with a plastic button base was tunneled through the incision, into the pocket, and out the

small opening, leaving the button under the skin. The button was sutured to the skin using

3-0 silk sutures (SofSilk, Tyco Healthcare Group Canada Inc., Ville St. Laurent, QC,

Canada), securing the externalized tether. To feed the catheter through the tether, the

catheter was first tunneled subcutaneously on the right side of the neck, from the ventral to

the dorsal incision areas using a 16G needle as a guide. The catheter was fed through the

underside of the plastic button and into the tether such that the end of the catheter

extended out of the tether. The tether and catheter were then secured to the tether-swivel

connector, which was fastened to a clamp on a retort stand. The catheter was tested again

to ensure for proper blood sampling and re-infusion. This rodent tethering system allowed

for manual, undisturbed blood sampling and for unrestricted movement of the rat. The

dorsal incision was closed with discontinuous 3-0 silk sutures. On the ventral side, the

catheter was anchored to the underside of the skin just cranial to the incision line, and the

incision was closed with continuous 3-0 silk sutures.

3.1.4.3 Post-operative care.

Following surgery, incision sites and the dorsal button area were cleaned with 10%

povidone-iodine. Rats were injected subcutaneously with 3.0 mL of saline to prevent

dehydration during recovery from anesthetization. Rats were then allowed to recover in

clean cages lined with paper towel overtop bedding material. The tether extended from the

back of the neck of the rat, through the cage lid, and connected to the swivel apparatus

external to the cage. Rats were allowed at least 3 full days to recover from surgery prior to

experimentation.

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3.1.4.4 Catheter patency and blood sampling.

To ensure that catheters remained free from clots, the lines were flushed daily with

heparinized (10 USP U/mL) saline. Carotid catheters were then filled with 100 USP U/mL

heparinized saline, plugged with a saline-filled syringe with a 23-gauge blunted needle, and

clamped with hemostats with silastic cuffs. Jugular catheters were filled with 100 USP

U/mL heparinized saline, plugged with blunted pins, and secured against the metal tether

with masking tape to avoid being caught or pulled.

3.2 General laboratory methods

3.2.1 Blood Glucose Determination

Whole blood glucose concentrations were measured immediately after obtaining blood

samples during experiments by the glucose oxidase method using a glucose analyzer

(Analox glucose analyzer, GMD-9D, Analox Instruments USA Inc., Lunenburg, MA). This

method was suitable for use with whole blood collected with heparin and/or EDTA. A 10 L

blood sample was pipetted into the reaction well, which contained the glucose oxidase

reagent (Glucose oxidase reagent kit, GMRD-020, Analox Instruments USA Inc., Lunenburg,

MA). In the presence of oxygen, glucose from the blood sample is oxidized by glucose

oxidase to gluconic acid and hydrogen peroxide as follows:

-D-glucose + O2  glucose oxidase

 D-gluconic acid + H2O2

Oxygen consumption is directly proportional to glucose concentration, and the analyzer

measures the rate of oxygen uptake in the reaction well using a Clark-type amperometric

oxygen electrode at 30C. The analyzer is calibrated before the start of, and intermittently

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throughout, each experiment using glucose standards of 8.0 (Glucose standard 8.0 mM,

GMRD-011) and 2.5 mM (Glucose standard 2.5 mM, RMRD-009), respectively. Results were

obtained within 35 sec of sample addition, and blood samples were analyzed in duplicate,

ensuring that the measurements obtained were ± 0.2 mM. The analyzer has a linearity of

30 mM and 50 mM using 10 L and 5 L samples, respectively.

For blood glucose measurements taken daily to monitor fed morning glycemia, a handheld

glucometer (Ascencia Elite, Bayer Canada Ltd., Etobicoke, ON, Canada) was used with blood

glucose test strips (Elite, Bayer Bayer Canada Ltd., Etobicoke, ON, Canada). Test strips

have been calibrated to give plasma equivalent glucose results. Test strips also use the

glucose oxidase method for determinations of glucose concentrations. The range of glucose

concentration measurable using test strips was 1.1 to 33.3 mM.

3.2.2 Preparation of blood sample collection tubes

1.5 mL microtubes (Eppendorf; Diamed Lab Supplies Inc., Mississuaga, ON, Canada) were

used to collect blood samples for various assays. Tubes chilled on ice containing 25 U of

dried heparin were used for collection of 0.1 mL of blood for corticosterone assays. For

glucagon, catecholamine, insulin, and growth hormone assays, 0.5 mL of blood was

collected in chilled tubes containing 50 L of a 1:1 solution of ethylenediamine tetraacetic

acid (EDTA; 0.24 g/mL in distilled water; Sangon Ltd. Canada, Scarborough, ON, Canada)

and Trasylol (2000 kallikrein IU; Bayer Canada Ltd., Etobicoke, ON, Canada). For

somatostatin assay, two tubes of 1.0 mL blood each were collected in chilled tubes each

containing 100 L of the 1:1 EDTA:Trasylol solution (i.e. 2.0 mL of blood collected in total at

euthanasia for somatostatin assay). EDTA is an anticoagulant, and Trasylol is a broad

spectrum proteolytic inhibitor that protected the plasma against loss of hormone reactivity.

These tubes were kept chilled (4C) prior to collection of blood.

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After blood was collected in the respective tubes, tubes were shaken to ensure adequate

mixing of their contents. Blood was then centrifuged (30-60 sec at 9000 rpm) to separate

plasma from packed red blood cells. Plasma was then immediately frozen and kept at -

80C.

3.2.3 Radioimmunoassays (RIA) for plasma hormone levels

Plasma corticosterone, glucagon, and catecholamine levels were measured using

radioimmunoassay (RIA). This procedure involves a competitive reaction between an

unknown amount of antigen (i.e. hormone) present in a plasma sample and a known

amount radiolabeled antigen for limited binding sites on an antibody (525). In our RIAs, the

radioactive label used was 125I. After an incubation period between these reagents during

which unlabeled and labeled hormones vie for a binding site on the antibody, an equilibrium

is reached. This equilibrium is dependent on the relative amounts of unlabeled and labeled

hormones and reflects the nature of competitive reactions. The bound hormone-antibody

complexes, which are in the form of a precipitant, are then separated from the free

hormones, which are in the liquid phase. This separation involves centrifuging the mixtures

and removing the liquid phase by decanting or aspirating. The amount of bound

radiolabeled hormone is then measured in a gamma scintillation counter. The assay is

calibrated by measuring standards of known concentrations and cross-plotting the counts

emitted by the radioactive label versus the concentration of the standards to generate a

dose-response curve. As the concentration increases, the number of counts decreases

exponentially. This is true also for the sample in question; the precipitated complex of a

sample carrying a high concentration of unlabeled hormone will have lower radioactivity

since there would be less radiolabeled hormone bound to the limited amount of antibody.

The reverse is also true in that a sample with a low concentration of unlabeled hormone will

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yield a precipitated pellet of lower radioactive signal. Non-specific binding is a control

parameter that represents the minimal binding (525). Percent bound values are calculated

for both standards and samples in the equation:

Percent bound = [ sample counts  ‐ non-specific binding counts ]

(zero calibrator counts  ‐ (non-specific binding counts)] × 100%

3.2.3.1 Plasma corticosterone assay.

ImmunoChem™ Double Antibody Corticosterone 125I RIA kits (MP Biomedicals, Solon, OH,

USA) were used to measure total corticosterone levels in rat plasma. Assay controls were

reconstituted with 2 mL of distilled water and allowed to sit at room temperature for at least

30 min before use. Other reagents from the kit and frozen plasma samples were also

brought to room temperature. Thawed samples were then centrifuged at 4C to remove

fibrin or other particulate matter. Plasma was diluted at a ratio of 1:200 by adding 2 mL of

steroid diluent to 10 L of plasma. The steroid diluent provided with the kit consisted of

phosphosaline gelatin buffer at pH 7.0 ± 0.1 with rabbit gamma globulins.

Six corticosterone standards in concentrations ranging from 0 to 1000 ng/mL were used to

produce a standard curve for the assay. 100 L of each of the standards, controls, and

diluted plasma samples were pipetted into polystyrene tubes. Standards were assayed in

triplicate while controls and samples were assayed in duplicate. To each of these tubes,

200 L of 125I-corticosterone followed by 200 L of anti-corticosterone antibody was added,

vortexed, and incubated at room temperature for 2 h. Corticosterone-3-

carboxymethyloxime:BSA was used as the antigen to generate antiserum in rabbits.

Following the incubation, 500 L of precipitating solution, a mixture of goat anti-rabbit

gamma globulin and polyethylene glycol in TRIS buffer, was added to the tubes. Tubes

66

were then vortexed and centrifuged at 2500 rpm for 20 min. This allowed for the

separation of bound antibody-antigen complexes (pellet) from the free fraction

(supernatant). A triplicate of tubes containing only 125I-corticosterone (to measure total

counts) and another triplicate of tubes containing only 125I-corticosterone and precipitating

solution (to measure non-specific binding) also underwent the same incubation and

vortexing procedures. The supernatant was then aspirated from all tubes, except for the

tube only containing 125I-corticosterone, and the precipitate was counted in a gamma-

scintillation counter to measure their reactivity. A standard curve was produced by plotting

percent bound against concentration of corticosterone for the standards. Plasma

concentrations of the samples were automatically calculated by computer software linked to

the gamma counter. The manufacturer’s assay precision was reported as intra-assay

coefficient of variation between 4.4-10.3% and inter-assay coefficient of variation between

6.5-7.1%.

3.2.3.2 Plasma glucagon assay.

Plasma glucagon levels were measured using LINCO/Millipore’s Glucagon RIA Kit (GL-32K,

LINCO Research Inc., St. Charles, MO, USA/Millipore Corporation, Billerica, MA, USA). This

assay uses 125-glucagon and glucagon antibody to determine the level of glucagon in plasma

by using the double antibody technique.

Samples were thawed just prior to the assay and centrifuged to separate plasma from fibrin

or other particulate matter. All reagents, besides the 125I-glucagon tracer, were provided

ready to use. 100 L of plasma was added with 100 L of assay buffer, which contained

0.2M glycine, 0.03M EDTA, 0.08% sodium azide, and 1% RIA-grade BSA at a pH 8.8 into

polystyrene tubes. When 100 L of plasma was not available, smaller volumes of sample

were used and additional assay buffer was added to compensate for the difference so that

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the total sample volume was equivalent to 100 L, as directed by the kit. Five standards

ranging from glucagon concentrations of 20 to 400 pg/mL were included in the kit. A zero

standard was produced using 200 L of assay buffer as instructed by the kit, and a 10

pg/mL standard was made by diluting the 20 pg/mL standard in a 1:1 ratio with assay

buffer. 100 L of each standard and control was pipetted into tubes along with 100 L of

assay buffer. To sample, standard, and control tubes, 100 L of guinea pig glucagon

antibody was added. All tubes were assayed in duplicate. Tubes containing 300 L of assay

buffer and no antibody were included in the assay to measure non-specific binding. Tubes

were vortexed, covered, and incubated at 4C for 20 hours.

Following the incubation period, lyophilized 125I-glucagon tracer was mixed with label

hydrating buffer, which had the same composition as assay buffer, and allowed to sit at

room temperature for 30 min. All tubes were then pipetted with 100 L of 125I-glucagon.

100 L of 125I-glucagon was added to two more tubes to measure total radioactivity (i.e.

total count tubes). All tubes were then vortexed, covered, and allowed to incubate at 4C

for an additional 20 hours.

After the second incubation period, 1.0 mL of cold (i.e. 4C) precipitating solution was added

to all tubes except for total count tubes. Tubes were then vortexed and incubated for 20

min at 4C after which they were centrifuged for 20 min at 4C. Supernatant was then

immediately aspirated from the tubes, except for total count tubes, and pellets containing

the antibody-antigen complexes were counted in a gamma-scintillation counter to measure

their radioactivity. A standard curve was plotted from the assay standards. Plasma

concentrations of the samples were automatically calculated by computer software linked to

the gamma counter. Samples which contained less than 100 L plasma were multiplied by

the appropriate factor. The intra-assay coefficient of variation was reported as 4.0-6.8%.

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The inter-assay coefficient of variation was reported as 7.3-13.5%. The specificity of the

test was reported as 100% for glucagon and <0.1% for oxyntomodulin, which contains the

glucagon sequence.

3.2.3.3 Plasma catecholamine assay.

A two-catecholamine RIA kit from Labor Diagnostika Nord (LDN) was used for the

quantitative determination of plasma epinephrine and norepinephrine (BA 1500, LDN GmbH

& Co. KG; Nordhorn, Germany). In this test, epinephrine and norepinephrine are first

extracted by using a cis-diol-specific affinity gel, acylated to N-acylepienphrine and N-

acylnorepienphrine, and then enzymatically converted during the detection procedure to N-

acylmetanephrine and N-acylnormetanephrine. The subsequent assay procedure follows the

basic RIA principle.

All reagents are brought to room temperature before use, except for the precipitating

solution which is kept chilled (~ 4C). Standards, controls, and plasma samples are first

extracted. Six standards ranging for each of epinephrine and norepinephrine are provided

with the kit in the concentration range of 0 to 90 ng/mL and 0 to 450 ng/mL, respectively.

20 L of each standard and controls and 80 L of distilled water are pipetted into wells of

the extraction plate, which is coated with boronate affinity gel. 100 L of sample plasma is

also pipetted into separate wells of the extraction plate. 50 L of each of assay buffer

(containing 1 M hydrochloric acid) and extraction buffer are added to all wells, and the plate

is covered and incubated at room temperature on an orbital shaker (600-900 rpm) for 30

min. Following the incubation period, standards, controls, and samples are acylated to N-

acylepinephrine and N-acylnorepinephrine. The plate is decanted, and 150 L of acylation

buffer followed by 25 L of acylation reagent are added to all wells. The plate is then

incubated under the same conditions as before for 15 min, decanted, pipetted with 1 mL of

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distilled water per well, incubated again for 5 min, and decanted again. 200 L of 0.025 M

hydrochloric acid is then added to each well, and the plate is covered and incubated as

before for 10 min. This eluate is then ready for RIA.

Epinephrine and norepinephrine RIAs are performed in separate tubes but from the same

extracted samples. Thus, the appropriate radioactively labeled antigen and antibody must

be used for each assay. 90 L of the extracted standards, controls, and samples are

pipetted into polystyrene tubes for each of the epinephrine and norepinephrine assays. To

measure non-specific binding, 90 L of hydrochloric acid is pipetted into separate tubes. An

enzyme provided with the kit containing catechol-O-methyltransferase is reconstituted in 1

mL of distilled water mixed thoroughly. To this solution, 0.3 mL of the coenzyme S-

adenosyl-L-methionine is then added, followed by 0.7 mL of the enzyme buffer. This

solution must be prepared no longer than 10 min prior to use. 25 L of enzyme solution is

added to all tubes. Tubes are then mixed and incubated at 37C for 30 min.

After the incubation, 50 L of 125I-epinephrine (or 125I-norepinephrine) is added to the

tubes. Tubes for measuring total radioactivity (i.e. total counts) are also included in the

assay and contain only the radioactive label. 50 L of epinephrine antibody (or

norepinephrine antibody) is then added to all tubes except tubes for non-specific binding or

total counts. After tubes are mixed and centrifuged at 500 xg and incubated at 4C

overnight. Cold precipitating reagent is mixed and 1 mL is then added to all tubes (except

total counts). Tubes are vortexed and centrifuged for 15 min at 3000 xg at 4C.

Supernatant is then aspirated, leaving behind a pellet containing antibody-antigen

complexes. Tubes for each assay were counted separately in a gamma-scintillation counter

to measure their radioactivity. A standard curve was plotted from the assay standards.

Plasma concentrations of the samples were automatically calculated by computer software

70

linked to the gamma counter. The intra-assay coefficient of variation was reported as 4.6-

14% and 4.0-4.6% for epinephrine and norepinephrine, respectively. The inter-assay

coefficient of variation was reported as 5.6-6.1% and 6.1-10% for epinephrine and

norepinephrine, respectively.

3.2.3.4 Plasma somatostatin assay.

The EURIA somatostatin RIA kit was used for the quantitative determination of plasma

somatostatin concentrations (RB-306, Euro-Diagnostica AB, Malmo, Sweden). Due to the

large volume of plasma needed for this assay, plasma was obtained from trunk blood at

euthanasia. Somatostatin from the plasma is first extracted with Sep-Pak C18 cartridges

(WAT023635, Waters Ltd., Milford, MA). The extracts are then assayed using 125-Tyr1-

somatostatin and an antibody to synthetic somatostatin-14 to determine the level of

somatostatin in the sample using the double antibody technique.

Plasma samples were thawed just prior to the assay and centrifuged to separate plasma

from fibrin or other particulate matter, and 0.5 mL of plasma was aliquoted into eppendorf

tubes. 50 L of 1M hydrochloric acid was added to each sample and vortexed. The Sep-Pak

cartridge was wetted with 5 mL of methanol and washed with 20 mL of distilled water.

Separate cartridges were used for each sample. At a flow rate no greater than 1 mL/10 sec,

the plasma-acid solution was applied to the Sep-Pak cartridge and then washed with 20 mL

of 4% acetic acid in distilled water. Somatostatin was subsequently eluted from the

cartridge with 2 mL of methanol also at a flow rate less than 1 mL/10 sec and collected in a

10 mL glass tube. The eluate was then dried in an evaporator. Extracted somatostatin

dissolved in 0.5 mL of assay diluents, which contained 0.05 M phosphate buffer at pH 7.4,

0.25% human serum albumin, 0.25% EDTA, 0.05% sodium azide, 0.1% Tween 80, and 500

71

KIU/mL Trasylol. Reconstituted samples were vortexed thoroughly and rested for 30

minutes prior to the RIA procedure.

For determination of the recovery in the extraction procedure, two recovery controls were

prepared. Control A consisted of 0.8 mL of rat plasma combined with 80 L of 1M

hydrochloric acid, to which 200 L of the 250 pmol/L somatostatin standard was added to

yield a final concentration of 50 pmol/L. Control B consisted of 0.8 mL of the same rat

plasma combined with 80 L of 1M hydrochloric acid, to which 200 L of assay diluents was

added. Both controls were extracted using the same procedure described above for the

samples. Control B was used for the correction for endogenous somatostatin in the

calculation of the recovery of added somatostatin.

A 250 pmol/L synthetic cyclic somatostatin (MW = 1638.1) standard was prepared in a

solution with same composition as the assay diluent and lyophilized. This lyophilized

standard was reconstituted in 5 mL distilled water and serially diluted with assay diluent to

yield 7 standard solutions ranging from 0 to 125 pmol/L. Rabbit somatostatin antibody was

raised against cyclic somatostatin conjugated to bovine thyroglobulin. The antibody and 125-

somatostatin were prepared in a solution with same composition as the assay diluent and

lyophilized, and both were reconstituted in 22 mL and 25 mL of distilled water, respectively,

prior to the RIA procedure. 100 L of standards, controls, and reconstituted samples were

pipetted into polystyrene tubes in duplicate. An additional pair of tubes with 300 L of

assay diluent was included to measure non-specific binding. 200 L of somatostatin

antibody was added to tubes containing standards, controls, and reconstituted samples and

vortexed. After a 20-h incubation at 4C, 200 L of 125- somatostatin was added to all tubes

and vortexed. Two additional tubes of 200 L of 125- somatostatin were included to measure

total count of radioactivity. After a second 20-h incubation at 4C, 100 L of precipitating

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reagent was added to all tubes except for total counts. This reagent consisted of rabbit

immunoglobulins coupled to cellulose particles and since it was a suspension, it required

continuous stirring during pipetting. Tubes were vortexed thoroughly, incubated for 45

minutes at 4C, and centrifuged at 2500 rpm at 4C for 20 min. Supernatant was then

immediately aspirated from the tubes, except for total count tubes, and pellets containing

the antibody-antigen complexes were counted in a gamma-scintillation counter to measure

their radioactivity. A standard curve was plotted from the assay standards. Plasma

concentrations of the samples were automatically calculated by computer software linked to

the gamma counter.

Somatostatin recovery was calculated as follows:

% recovered somatostatin = [ concentration Control A  – concentration Control B ]

50 × 100%

The recovery should be at least 60% for a valid assay. Sample concentrations were then

corrected for the percent recovery and also multiplied by a factor of 1.1 to correct for the

increased sample volume by adding 1M hydrochloric acid. The intra-assay coefficient of

variation was reported as 2.8-8.3%. The inter-assay coefficient of variation was reported as

3.3-6.4%.

3.2.3.5 Plasma insulin assay

Plasma insulin levels were measured using LINCO/Millipore’s Rat Insulin RIA Kit (RI-13K,

LINCO Research Inc., St. Charles, MO, USA/Millipore Corporation, Billerica, MA, USA). This

assay uses 125-insulin and a rat insulin antibody to determine the level of rat insulin in

plasma by using the double antibody technique.

73

Samples were thawed just prior to the assay and centrifuged to separate plasma from fibrin

or other particulate matter. Since high doses of insulin were administered in the

hypoglycemia experiments, plasma samples obtained after insulin administration had to be

diluted with assay buffer before assay. For Study 1, when 10 U of insulin was injected,

plasma samples at time 60 and 180 min were diluted 100 and 20 times, respectively. For

Study 2, plasma samples at time 60 were diluted 100 times.

All reagents, besides the 125I-rat insulin tracer, were provided ready to use. One-half of the

volumes of all reagents suggested in the manufacturer’s protocol were used in this assay.

50 L of plasma (or diluted plasma) was pipetted into glass sample tubes. 100 L and 50 L

of assay buffer, which contained 0.05 M phosphosaline at pH 7.4, 0.025M EDTA, 0.08%

sodium azide, and 1% RIA-grade BSA, was added to the non-specific binding tubes and

reference tubes, respectively. Seven standards ranging from rat insulin concentrations of

0.1 to 10.0 ng/mL were included in the kit. 50 L of each standard and control was pipetted

into tubes. All tubes were assayed in duplicate. Lyophilized 125I-insulin tracer was hydrated

with 27 mL of hydrating buffer and rested at room temperature for 30 min with occasional

gentle mixing. The hydration buffer contained normal guinea pig immunoglobulin G. 50 L

of hydrated 125I-insulin was added to all tubes. 50 L of 125I-insulin was added to two more

tubes to measure total radioactivity (i.e. total count tubes). To sample, standard, and

control tubes, 50 L of guinea pig rat insulin antibody was added. The rat insulin antibody

was raised in guinea pigs against highly purified rat insulin. All tubes were assayed in

duplicate. Tubes were vortexed, covered, and incubated at 4C for 20 hours.

After the incubation period, 0.5 mL of cold (i.e. 4C) precipitating solution was added to all

tubes except for total count tubes. The precipitating solution contained goat anti-guinea pig

immunoglobulin G serum, 3% polyethylene glycol, 0.05% TritonX-100 in 0.05 M

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phosphosaline, 0.025 M EDTA, and 0.08% sodium azide. Tubes were then vortexed and

incubated for 20 min at 4C after which they were centrifuged for 20 min at 4C.

Supernatant was then immediately aspirated from the tubes, except for total count tubes,

and pellets containing the antibody-antigen complexes were counted in a gamma-

scintillation counter to measure their radioactivity. A standard curve was plotted from the

assay standards. Plasma concentrations of the samples were automatically calculated by

computer software linked to the gamma counter. Samples which were diluted were

multiplied by the appropriate factor. The intra-assay coefficient of variation was reported as

1.4-1.6%. The inter-assay coefficient of variation was reported as 8.5-9.4%. The

specificity of the test was reported as 100% for rat, human, and porcine insulin and not

detectable for IGF-I or rat C-peptide.

3.2.3.6 Plasma growth hormone assay

Plasma growth hormone levels were measured using LINCO’s Rat Growth Hormone RIA Kit

(RGH-45HK, LINCO Research Inc., St. Charles, MO, USA). This assay uses 125-rat growth

hormone and a rat growth hormone antibody to determine the level of rat growth hormone

in plasma by using the double antibody technique.

Samples were thawed just prior to the assay and centrifuged to separate plasma from fibrin

or other particulate matter. All reagents, besides the 125I-growth hormone tracer, were

provided ready to use. 100 L of plasma was pipetted into sample tubes containing 100 L

assay buffer. The assay buffer contained 0.05 M phosphosaline at pH 7.4, 0.025M EDTA,

0.08% sodium azide, and 1% RIA-grade BSA. 300 L and 200 L of assay buffer was added

to the non-specific binding tubes and reference tubes, respectively. Seven standards

ranging from rat growth hormone concentrations of 0.5 to 50.0 ng/mL were included in the

kit. 100 L of each standard and control was pipetted into tubes. All tubes were assayed in

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duplicate. Lyophilized 125I-growth hormone tracer was hydrated with 13.5 mL of hydrating

buffer and rested at room temperature for 30 min with occasional gentle mixing. The

hydration buffer contained normal guinea pig serum. 100 L of hydrated 125I-growth

hormone was added to all tubes. 100 L of 125I-growth hormone was added to two more

tubes to measure total radioactivity (i.e. total count tubes). Both the 125I tracer and

standards were prepared with recombinant rat growth hormone. To sample, standard, and

control tubes, 100 L of guinea pig rat growth hormone antibody was added. The rat

growth hormone antibody was raised in guinea pigs against recombinant rat growth

hormone. All tubes were assayed in duplicate. Tubes were vortexed, covered, and

incubated at 4C for 20 hours.

After the incubation period, 1.0 mL of cold (i.e. 4C) precipitating solution was added to all

tubes except for total count tubes. The precipitating solution contained goat anti-guinea pig

immunoglobulin G serum, 3% polyethylene glycol, 0.05% TritonX-100 in 0.05 M

phosphosaline, 0.025 M EDTA, and 0.08% sodium azide. Tubes were then vortexed and

incubated for 20 min at 4C after which they were centrifuged for 20 min at 4C.

Supernatant was then immediately aspirated from the tubes, except for total count tubes,

and pellets containing the antibody-antigen complexes were counted in a gamma-

scintillation counter to measure their radioactivity. A standard curve was plotted from the

assay standards. Plasma concentrations of the samples were automatically calculated by

computer software linked to the gamma counter. The specificity of the test was reported as

100% for rat growth hormone and <0.1% for rat prolactin and human and porcine growth

hormone.

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3.2.4 Pancreatic protein determination

Pancreatic glucagon and somatostatin protein content was measured from pancreata

collected from animals in Study 1. Pancreata were quickly removed at euthanasia, frozen in

sterile tubes on dry ice, and stored at -80°C.

3.2.4.1 Pancreas homogenization for protein isolation

Frozen pancreata were individually homogenized in 3 10-second intervals on medium-high

speed in 50-mL falcon tubes kept on ice in 10 mL of extraction medium containing 9.2%

hydrochloric acid, 5% formic acid, 1% trifluoroacetic acid (TFA), and 1% sodium chloride.

Homogenates were centrifuged at 2300 rpm at 4C for 15 minutes. The supernatant was

collected in a separate falcon tube. An additional 10 mL of extraction medium was added to

the remaining pellet, re-homogenized as before, and centrifuged again at 2300 rpm at 4C

for 15 minutes. The second supernatant was collected in the same tube as the first (total

final volume of 20 mL), and the pellet was discarded.

3.2.4.2 Sep-Pak protein extraction

Pancreatic protein was isolated by eluting the supernatant of pancreatic homogenates using

Sep-Pak C18 cartridges (WAT023635 Waters Ltd., Milford, MA). Separate cartridges were

used for each sample. Using a 20 mL syringe, each Sep-Pak cartridge was first wetted with

15 mL of a solution containing 80% isopropanol and 0.1% TFA, followed by 15 mL of 0.1%

TFA. The 20 mL of supernatant from the homogenized pancreata were then slowly passed

through the Sep-Pak twice. Each Sep-Pak cartridge is subsequently washed using 15 mL of

0.1% TFA to remove any unbound sample. At the end of this stage only is air allowed to

pass through the column to remove all liquid (approximately 2 mL of air from a syringe).

The protein is then eluted from the column using 10 mL of solution containing 80%

isopropanol and 0.1% TFA into a 15 mL falcon tube. Pancreatic protein eluates were frozen

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at -80°C until used. For glucagon and somatostatin determinations using commercially

available radioimmunoassay kits, 1-mL aliquots of these eluates were evaporated, and

isolated protein was re-dissolved in 3 mL of double-distilled water. Glucagon and

somatostatin measurements were normalized for total protein content as determined by

spectrophotometer analysis.

3.2.5 Liver mRNA expression quantification

Livers were quickly removed at euthanasia, frozen in sterile tubes on dry ice, and stored at -

80°C.

3.2.5.1 Total RNA preparation.

Total RNA was extracted from frozen liver using a modified phenol-chloroform method with

TRI Reagent® (TR-118, Sigma-Aldrich, St. Louis, MO, USA). TRI Reagent® combines phenol

and guanidine thiocyanate to facilitate inhibition of RNAse activity. A small sample of liver

(approximately 2 x 2 x 2 mm3) was homogenized in a 15 mL snap cap tube for 20-30 on

medium-high speed in a fume hood. The homogenate was then incubated at room

temperature for at least 5 min. 800 L of chloroform was then added to the homogenate

and shaken vigorously before incubating for an additional 5 min at room temperature.

Using a 1 mL pipette, the upper phase (approximately 2 mL of clear fluid) of the mixture

was carefully collected in a new 15 mL snap cap tube with care being taken to avoid the

buffy white layer to avoid DNA contamination. At this stage, RNA remains exclusively in the

aqueous phase (supernatant), DNA in the interphase (buffy white layer), and proteins in the

organic phase (pellet). 2 mL of isopropanol was then added to the supernatant collected,

and the tube was mixed by inversion and incubated at room temperature for 15 min. Tubes

containing the samples were centrifuged at 12 000 g at 4C for 10 min. The orientation of

the tubes was noted prior to centrifugation so that the small RNA pellet could be easily

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identified following centrifugation. The supernatant was then removed by aspiration, and 4

mL of ice-cold 75% ethanol was added to the remaining pellet. Tubes were then vortexed

thoroughly and centrifuged at 10 000 g at 4C for 5 min. The supernatant was then

carefully removed by aspiration. Tubes were then allowed to vent at room temperature for

30 min to evaporate off any remaining ethanol. Pellets were resuspended in 100 L

diethylpyrocarbonate (DEPC) water (i.e. RNAse free water) by repeated drawing and

expelling the pellet with a pipette tip.

Quantification of RNA content was performed by measuring the optical density (OD) at 260

and 280 nm using 2 L of sample with a NanoDrop 1000 spectrophotometer (Thermo Fisher

Scientific, Mississauga, ON, Canada). For RNA, the ratio of 260/280 should be between 1.8

and 2. OD measurements below this ratio are indicative of contamination with protein or

solvents. RNA concentration (in g/mL) was then calculated as:

RNA concentration = OD260 × dilution factor × 40

The resuspended RNA sample was transferred to eppendorf tubes and stored at -80C until

use.

3.2.5.2 cDNA synthesis.

Frozen RNA dissolved in DEPC water was heated in a water bath at 65C for 5 min. Using

concentrations obtained from spectrophotometer determinations, 10 g of RNA was very

gently combined with 5 L DEPC water, 2 L DNAse buffer (10x), and 3 L DNAse in sterile

microtubes. After a 5-min incubation at room temperature, the DNAse reaction was

stopped with incubation with 2 L 25mM EDTA at 70C for 12 min.

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Following treatment with DNase I, purified RNA was reverse transcribed to synthesize first-

strand cDNA using SuperScript™III (Invitrogen Corporation, Carlsbad, CA, USA). 2 L of

random primers (12-times dilution in DEPC water; 48190-011, Invitrogen Corporation) 2 L

10 mM dNTP (Fermentas, Thermo Fisher Scientific, Mississauga, ON, Canada) were added to

the DNAse-treated RNA and heated at 65C for 5 min and cooled on ice for 1 min. 8 L

First-Strand buffer (5x, Invitrogen Corporation) and 2 L 0.1 M DTT (Invitrogen

Corporation) were added to the tubes. The contents of each tube (total volume = 36 L)

was then equally divided into 2 sets of tubes (18 L per tube): RT+ tubes to which 1 L of

SuperScript™III Reverse Transcriptase (Invitrogen Corporation) was added and RT- tubes

(a negative control for reverse transcription), to which 1 L DEPC water was added. The

tubes were then incubated at 25C for 10 min (reaction for random primers), 50C for 60

min (reverse transcription reaction), and 70C for 15 min (inactivation of reaction) in a

thermal cycler (Vapo Protect Mastercycler® pro, Eppendorf Canada, Mississauga, ON,

Canada). RNA complementary to the cDNA was then removed by adding 1L RNAse H (2.5-

times dilution in RNAse buffer; Fermentas) and incubating at 37C for 20 min. The cDNA

can then be used as a template for amplification.

3.2.5.3 Polymerase chain reaction (PCR) of -actin.

This PCR was used to determine the integrity of the RT+ and RT- sets of synthesized cDNA.

Samples from RT+ tubes should demonstrate amplification of the DNA of interest (-actin),

whereas RT- tubes which lacked reverse transcriptase enzyme should not amplify DNA.

Signals that appear in the RT- tubes are indicative of DNA contamination. A PCR mixture is

first made for all samples to be run by combining the following reagents using the following

ratio per 1 L cDNA sample: 11 L double-distilled water, 0.25 L 5’ (forward) primer for -

actin (ACGT Corp, Toronto, ON, Canada), 0.25 L 3’ (reverse) primer for -actin (ACGT

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Corp), and 12.5 L REDTaq® (Sigma-Aldrich, St. Louis, MO, USA). Care should be taken to

gently combine the REDTaq® solution, which contains the enzyme Taq DNA polymerase. In

96-welled plates, 1 L of cDNA from each of RT+ and RT- sets of tubes was first pipetted

before 24 L of the PCR mixture was then added to each well. Plates were then spun at

1000 rpm at 4C for 30 sec. After thermal cycler PCR machine was warmed up, plates were

sealed with a film to prevent evaporation and placed in the thermal cycler at 94C for 5 min

(initial denaturation) and cycled 40 times at 94C for 30 sec, 45C for 30 sec, and 72C for

30 sec (amplification) prior to incubation at 72C for 4 min (final elongation), and then 4C

for storage.

A 1% agarose running gel for -actin was prepared by combining 200 mL 1x TBE (Tris, boric

acid, and EDTA), 2 g agarose powder, and 20 L SYBR® Safe (10 000x, Invitrogen

Corporation) DNA gel stain, heating until completely dissolved, and setting into a 20-comb

cast. The first well of each row was reserved for 10 L of DNA ladder (GeneRuler™, 1 kb,

Fermentas), and 10 L of sample was loaded into the wells. Charge was applied to the gel

at approximately 90 V, and samples travelled through the gel from the negative to positive

electrode. Blots on finished gels were visualized under ultra violet light in a UVP BioDoc-It

System (UVP, Upland, CA, USA). Real-time quantitative PCR was run only for samples that

showed no contamination.

3.2.5.4 Real-time quantitative PCR.

Real-time quantitative PCR was run only for samples that showed no contamination.

Several RT- samples and a DEPC-water sample were included in the run to serve as

controls. All samples were run in duplicate. 1 L of cDNA and 4 L of yellow-coloured DEPC

water was pipette into wells of MicroAmp® Optical 384-Well Reaction Plates (Applied

Biosystems, Foster City, CA, USA). The yellow colouring allowed for easier detection when

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working with such small volumes. For cyclophilin, cDNA was first diluted 50 times prior to

adding 1 L of the diluted cDNA to the wells. Real-time quantitative PCR was performed for

using primers for 2 genes of interest (PEPCK1 (Pck1), Mm01247058_m1; G6Pase (G6pc),

Mm00839363_m1) and 1 housekeeping gene (cyclophilin (Ppia), Mm02342430_g1) using

TaqMan Gene Expression Assays (Applied Biosystems). A reaction mixture of TaqMan®

Universal PCR Master Mix (Applied Biosystems), primer of the gene, and blue-coloured DEPC

water was combined together in a ratio of 10:1:4 prior to pipetting 15 L of this mixture

into each well. Plates were sealed with MicroAmp® Optical Adehesive Film (Applied

Biosystems) and spun at 1000 for 1 min at 4C.

Real-time quantitative PCR was performed with ABI Prism 7900 Sequence Detection System

with Sequence Detection System v2.1 software (Applied Biosystems). The thermal cycling

conditions for the reaction consisted of 2 min at 50C, 10 min at 95C, and 40 cycles of 15

sec at 95C and 1 min at 60C. After plates were run, threshold cycle (Ct) values were

analyzed. Ct values should be between 20-25; cDNA needs to be diluted if Ct values are

below this range. If Ct values fall above this range, cDNA needs to be more concentrated,

or optimization of the primer is necessary. Ct values for the samples run were within the

optimal range. The expression values of each gene of interest were normalized to the

internal control gene, cyclophilin, for the efficiency of amplification and quantified by the 2-

Ct method (526). Cyclophilin expression values did not vary between experimental groups,

deeming it a suitable control gene.

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3.3 Preparation of SSTR2 antagonist and saline infusates

This peptide (referred to in literature by the names PRL-2903, DC-41-33, and BIM-23458)

was synthesized and provided by Dr. David Coy (Tulane University, New Orleans, LA, USA)

in lyophilized form. The peptide was stored at -20C in a jar of desiccant. To prepare

stock-solutions for long-term storage, the peptide was carefully weighed and aliquoted into

eppendorf tubes, and dissolved in a calculated volume of 1% acetic acid to a concentration

of approximately 25 000 nmol/mL. Each vial of aliquot was carefully labeled for its

concentration, sealed with parafilm, and stored also at -20C. Repeated thawing and

refreezing was avoided. On the morning of the experiment, a vial of aliquoted peptide stock

solution was thawed on ice and diluted with 0.9% saline (NaCl solution)specific to the dose

used for each experiment (1500 nmol/kg/h in Study 1 or 3000 nmol/kg/h in Study 1 and

Study 2). In Study 1, 5 mL of infusate was prepared per animal, and in Study 2, 6 mL of

infusate was prepared per animal. This volume was used to allow for enough infusate for

the 4 or 5 h infusion period for each of Study 1 and Study 2, respectively, at an infusion

rate of 1 mL/h in addition to the volume needed to prime the infusion lines. Infusates were

neutral in pH. Rats that did not receive SSTR2 antagonist were given infusions of vehicle

solution (i.e. 1% acetic acid in 0.9% saline at a ratio of approximately 250 L : 5 mL).

3.4 Statistical analysis

Statistical analysis was performed using Statistica software (Statsoft Inc., Tulsa, OK, USA).

In all tests, significance was deemed when P < 0.05. Unless otherwise stated, all data are

expressed as mean ± standard error of the mean (SEM). Outlying data points were

removed when they occurred greater than two standard deviations from the mean.

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One-Way Analysis of Variance (ANOVA) was used to compare metabolic parameters, basal

hormone values, hormone responses, areas under the curve (AUC), and tissue mRNA and

protein levels between groups, unless otherwise stated. AUC was calculated using Prism

software (GraphPad Software, San Diego, CA, USA). Measurements that were taken

repeatedly over time were compared using Repeated Measures ANOVA. If differences

between groups were significant using ANOVA, Duncan’s post-hoc test was used to

determine which groups differed from each other. In Study 2, comparisons within the same

group on Experimental Day 1 and Experimental Day 2 were assessed using a paired T-test.

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84

Chapter 4 Study 1: Effect of SSTR2 antagonism on

counterregulation to hypoglycemia

4 Study 1

4.1 Aims

The objective of this study was to determine whether the glucagon response to

hypoglycemia can be improved or normalized in diabetic rats by removing the suppressive

effect of somatostatin on glucagon secretion. Different subtypes of the somatostatin

receptor within the pancreatic islet mediate the inhibitory effects of somatostatin on islet

hormone secretion. In rodent and human islets, somatostatin receptor type 2 (SSTR2) are

found nearly exclusively on glucagon-secreting alpha-cells while SSTR1 and SSTR5 are

abundant on human and human and rodent beta-cells, respectively (385;389). In this

study, we use an antagonist to SSTR2, via which somatostatin exerts its inhibitory effect on

stimulated glucagon secretion. Using this selective antagonist to SSTR2, Efendic and

colleagues demonstrated that stimulated-secretion of glucagon is dose-dependently

enhanced in perifused islets and perfused pancreata of non-diabetic rats. Increasing

somatostatin concentrations dose-dependently reverses the effectiveness of the SSTR2

antagonist, which demonstrates a competitive nature for the SSTR2 (392). We hypothesize

that in diabetes during hypoglycemia, glucagon release is, at least in part, suppressed by

the prevailing inhibitory effect of excessive somatostatin acting via SSTR2. Thus, we

hypothesize that selective SSTR2 antagonism will normalize the glucagon response to

hypoglycemia.

85

Somatostatin also inhibits the release of other counterregulatory hormones: the

hypothalamic-pituitary-adrenal (HPA) axis (corticotrophin-releasing hormone (CRH) and

adrenocorticotrophic hormone (ACTH)) and the sympathoadrenal system (epinephrine and

norepinephrine) (271). Intracerebroventricular administration of somatostatin or its

analogues decreases stress-induced glucocorticoid and catecholamine responsiveness

(169;527;528). Thus, we also evaluated the effect of SSTR2 antagonism on other

counterregulatory hormone responses during hypoglycemia. Presently, we demonstrate

that SSTR2 antagonism restores glucagon and corticosterone counterregulation to

hypoglycemia in diabetic rats. In the absence of hypoglycemia, this SSTR2 antagonist

neither elicits hyperglycemia nor markedly elevates counterregulatory hormone levels.

Finally, we wished to determine whether improved glucagon and corticosterone

counterregulation could: 1) reduce the potential risk of hypoglycemia in an unclamped

condition as would be indicated by a decrease in the exogenous glucose requirement during

our conditions of hypoglycemic clamp; and 2) increase the mRNA expression of

gluconeogenic enzymes, phosphoenolpyruvate carboxykinase (PEPCK) and glucose-6-

phosphatase (G6Pase), despite the large dose of insulin necessary to induce hypoglycemia

in our diabetic rats.

4.2 Experimental methods

4.2.1 Study design

The experimental protocol of Study 1 is shown in Figure 4-1. Three main groups of rats –

non-diabetic controls (N), diabetic controls (D), and diabetic rats given somatostatin

receptor type 2 antagonist (D+SSTR2a) – were used in hypoglycemia clamp experiments

and in control experiments during basal conditions (i.e. in the absence of exogenously

86

administered insulin). An additional group to assess the effects of SSTR2a during

hypoglycemia in non-diabetic rats (N+SSTR2a) was also included. Schematics to illustrate

the procedures for hypoglycemia clamp and control experiments are shown in Figure 4-2A

and Figure 4-2B, respectively.

4.2.2 Hypoglycemia experiments

After basal samples were obtained at t=-60 min, infusions of saline or SSTR2 antagonist

(3000 nmol/kg/h) at 1 mL/h were started for controls and D+SSTR2a rats, respectively.

Infusions were delivered via digital pumps (Harvard Apparatus PHD 22/2000 syringe pumps,

Holliston, MA). A SSTR2 antagonist dose of 1500 nmol/kg/h was initially tested in our pilot

studies based on unrelated in vivo studies in non-diabetic rats using the same antagonist

(508). However, since the effect of the SSTR2 antagonist at 1500 nmol/kg/h to enhance

glucagon release during hypoglycemia was only moderate in diabetic rats, we rationalized

that a larger dose of the antagonist may be necessitated since diabetic rats have increased

somatostatin. Blood glucose was measured a glucose analyzer in duplicate at the time

points -60, -40, -20, 0, 5, 10, 15, 20, and every 10 min thereafter until 180 min. Blood

samples were obtained from the carotid catheter at regular time intervals throughout the

glucose clamp and from trunk blood at euthanasia for measurement of plasma hormone

levels. After plasma was removed, packed red blood cells were re-suspended in heparinized

saline (10 USP U/mL) containing 1% bovine serum albumin and re-infused into the rat.

After blood samples were obtained at t=0, regular insulin (10 U/kg; Humulin R, Eli Lilly,

Indianapolis, IN) was injected subcutaneously in all rats (N, n=14; D, n=14, D+SSTR2a,

n=18). In this study, we chose to give a subcutaneous injection of insulin rather than

insulin infusion to induce hypoglycemia. This method of insulin administration is more

similar to the clinical insulin regime where hypoglycemia is more relevant. Also, we have

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noticed a more marked impairment of the glucagon response to hypoglycemia using insulin

injection (174) than insulin infusion (529), despite a similar amount of insulin administered,

and we wished to test the efficacy of our SSTR2 antagonist in a setting of greater glucagon

unresponsiveness. However, this method of insulin bolus rather than infusion does not

result in a steady state of insulin action, and therefore it makes it difficult to measure

glucose turnover precisely, which is why we did not use glucose tracer techniques. To

evaluate the effect of a lower insulin dose with the same SSTR2 antagonist dose (3000

nmol/kg/h) on glucose infusion rates, hypoglycemia was also induced using 5 U/kg (N, n=5;

D, n=4, D+SSTR2a, n=4). Glucose infusions (50% dextrose) were given at a variable rate

to clamp glycemia at a target hypoglycemia of 2.5±0.5 mM. After t=180 min, rats were

quickly euthanized by decapitation. Plasma from trunk blood and organs were collected,

frozen immediately, and stored at -80°C until assayed.

4.2.3 Control experiments: SSTR2 antagonist in the absence of hypoglycemia

To evaluate the effect of SSTR2 antagonist infusion per se, glycemia and plasma glucagon,

corticosterone, and catecholamines were measured in the absence of hypoglycemia in a

separate set of non-diabetic (Ctrl:N, n=7), diabetic (Ctrl:D, n=7), and diabetic rats given

somatostatin receptor type 2 antagonist (Ctrl:D+SSTR2a, n=7) under basal conditions for

240 min. The same methodology and sampling procedure was used during these control

experiments with the following exceptions: saline rather than insulin was injected at t=0,

and glucose infusions were not used. SSTR2 antagonist was infused at 3000 nmol/kg/h in

these control experiments (Ctrl:D+SSTR2a).

4.2.4 Effect of SSTR2 antagonist in non-diabetic rats during hypoglycemia

The effect of SSTR2 antagonism in non-diabetic rats was also assessed during

hypoglycemia. This control group allows (N+SSTR2a) for an assessment of the diabetic

88

impairment that might be attributable to elevated somatostatin. Using the same protocol as

the hypoglycemia experiments described above, glucagon and corticosterone responses to

hypoglycemia in non-diabetic rats under conditions of high insulin dose (10 U/kg) and 1500

(n=3) and 3000 (n=4) nmol/kg/h SSTR2 antagonist and at low insulin dose (5 U/kg) and

3000 nmol/kg/h (n=10) were assessed and compared to the corresponding non-diabetic

controls (N) described above.

4.3 Results

4.3.1 Daily blood glucose, body weight, and food intake

Metabolic parameters such as fed morning blood glucose, body weight, and food intake

were measured daily to monitor the health of all rats and to ensure that diabetic rats were

of a similar diabetic state leading up to the experimental day before being randomly

assigned into control or SSTR2 antagonist treatment groups. Fed morning glycemia

significantly increased 4-fold in D compared with N in all three hypoglycemia (including the

pilot study) (P<0.0002) and control (P<0.0002) studies (Table 4-1). Diabetic groups within

each study had similar hyperglycemia leading up to the clamps. Similarly, D had

significantly decreased body weight as compared to N in all three hypoglycemia (P<0.03)

and control (P<0.03) studies and similar body weight to D+SSTR2a (Table 4-1). Average

daily food intake also significantly increased in D compared with N in all hypoglycemia

(P<0.0002) and control (P<0.0001) studies (Table 4-1). D+SSTR2a rats in the

hypoglycemia study group with the dosing combination of 10 U/kg insulin and 3000

nmol/kg/h had a slightly lower food intake (P<0.04) as compared to D, despite having

similar basal glycemia and body weight. The metabolic changes associated with 3-weeks of

diabetes (i.e. severe postprandial hyperglycemia and decreased body weight compared to

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non-diabetic controls despite hyperphagia) have been well-documented and represent a

slew of physiologic processes which, taken together, can be summarized as accelerated

catabolism. Simply stated, the metabolic derangements observed in diabetic subjects result

in large from a state of relative insulin deficiency, which leads to catabolic processes

including gluconeogenesis, lipolysis, ketogenesis, and proteolysis (530).

4.3.2 Hypoglycemia clamp experiments

4.3.2.1 Pilot study: using 10 U/kg insulin and 1500 nmol/kg/h SSTR2 antagonist.

In our pilot studies, we initially used SSTR2 antagonist at a dose of 1500 nmol/kg/h, a dose

based on previous in vivo studies in non-diabetic rats (508). SSTR2 antagonist at a dose of

1500 nmol/kg/h in our initial pilot studies yielded significant but very modest improvements

of the glucagon response (Figure 4-3A) and modest and insignificant increases of

corticosterone (Figure 4-3B). We speculated that a larger dose of SSTR2 antagonist would

be required to overcome the increased amounts of pancreatic and circulating somatostatin

observed in STZ-diabetic rats and doubled the dose of SSTR2 antagonist in subsequent

experiments.

4.3.2.2 Using 10 U/kg insulin and 3000 nmol/kg/h SSTR2 antagonist.

D had elevated basal plasma glucagon (P<0.004), corticosterone (P<0.006), and

epinephrine (P<0.04) levels as compared with N, and both groups of diabetic rats had

similar basal plasma hormone levels before administration of SSTR2 antagonist (Table 4-2).

Basal plasma insulin in D was reduced to one-half of the levels in N (P<0.05) and similar in

both diabetic groups (Table 4-3).

In the hypoglycemia studies, all groups reached target hypoglycemia (2.5 ± 0.5 mM) by 70

min after insulin injection and remained there until the end of experiment at 180 min

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(Figure 4-4). N rats reached target hypoglycemia at time 50 min slightly before the diabetic

groups.

As expected, N exhibited a robust, 11-fold increase in plasma glucagon whereas D had a

markedly attenuated glucagon response (P<0.0002 vs N). The glucagon response in

D+SSTR2a was fully normalized (P<0.0001 vs D; Figure 4-5A). Similarly, the

corticosterone response was significantly blunted in D as compared to the 10-fold increase

observed in N (P<0.002) but was greatly improved in D+SSTR2a (P<0.05 vs D; Figure

4-5B).

In the high-dose insulin hypoglycemia clamps, glucose infusion rates (mean infusion rates

N: 8.1±1.0; D: 8.9±1.2; D+SSTR2a: 9.2±1.2 mg/kg/min) nor the total amount of glucose

infused to maintain target hypoglycemia (Figure 4-5C) differed amongst groups.

N rats exhibited a 69-fold increase in plasma epinephrine, and interestingly, D rats attained

a similar plasma epinephrine level to N rats, suggesting that in this model of hypoglycemia,

diabetic rats do not exhibit a defect of epinephrine counterregulation (Figure 4-6A). SSTR2

antagonist administration did not alter the epinephrine response to hypoglycemia (Figure

4-6A), presumably because the maximal epinephrine response was already achieved.

Norepinephrine levels were likewise similar in all groups, achieving 3- to 5-fold increases in

responses to hypoglycemia (Figure 4-6B).

Plasma insulin levels after high-dose (10 U/kg) insulin injection increased approximately

144- to 200-fold in all three groups due to the large amount of insulin required to induce

hypoglycemia in diabetic rats which are otherwise quite insulin resistant (Table 4-3). The

same dose was used in non-diabetic rats to allow for more relevant comparisons with

diabetic groups.

91

4.3.2.3 Using 5 U/kg insulin and 3000 nmol/kg/h SSTR2 antagonist.

An overnight fast lowered initial starting blood glucose levels in both diabetic groups to

similar euglycemic levels as non-diabetic rats, despite marked fed glycemia (Table 4-1) and

basal plasma insulin at half the concentration of non-diabetic rats (Table 4-3). In the low

dose insulin hypoglycemia studies, all groups reached target hypoglycemia (2.5 ± 0.5 mM)

30 min after insulin injection and remained there until the end of experiment at 180 min

(Figure 4-7).

Basal plasma glucagon levels were increased in both diabetic groups as compared with non-

diabetic rats (Table 4-3). However, basal plasma corticosterone, epinephrine, and

norepinephrine were similar in all groups (Table 4-3). D rats also demonstrated markedly

attenuated glucagon (P<0.04; Figure 4-8A) and corticosterone (P<0.04; Figure 4-8B)

responses to hypoglycemia induced with a lower dose (5 U/kg) of insulin. D+SSTR2

exhibited fully restored glucagon (P<0.005; Figure 4-8A) and corticosterone (P<0.04;

Figure 4-8B) responses. Thus, the SSTR2 antagonist was effective in improving glucagon

and corticosterone counterregulation to hypoglycemia induced by both the high and low

doses of insulin tested.

In the low-dose insulin hypoglycemia clamps, D+SSTR2a required lower glucose infusion

rates (mean infusion rates: N: 14.1±1.4; D: 15.5±1.5; D+SSTR2a: 6.2±1.2 mg/kg/min;

P<0.001, D vs D+SSTR2a) and had a lower total glucose requirement than D (P<0.008;

Figure 4-8C).

N exhibited an 87-fold increase in plasma epinephrine, and interestingly, D attained

epinephrine levels similar to N, suggesting that 3-week STZ-diabetic rats do not exhibit a

defect of epinephrine counterregulation (Figure 4-9A). Norepinephrine levels were likewise

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similar in all groups, achieving 2- to 3-fold increases in response to hypoglycemia (Figure

4-9B).

Plasma insulin levels after low-dose (5 U/kg) insulin injection increased to approximately 60

ng/mL in N, D, and D+SSTR2a (Table 4-3).

Basal plasma growth hormone levels were significantly lower in D as compared to N

(P<0.01), and both diabetic groups had similar basal plasma growth hormone levels (Figure

4-10). Plasma growth hormone levels were similar in all groups during hypoglycemia.

Notably, administration of SSTR2 antagonist in diabetic rats did not affect growth hormone

concentrations.

4.3.3 Effect of SSTR2 antagonist in the absence of hypoglycemia

All groups that underwent control experiments had similar basal hormone levels (Table

4-4). The lack of difference in glucagon and corticosterone between Ctrl:N and Ctrl:D may

be attributed to elevated levels of these basal hormones in Ctrl:N, which suggests that

Ctrl:N rats may have been a mildly stressed despite the extreme care taken to avoid stress

during handling.

Glycemia remained steady throughout the 4-h experiment (Figure 4-11). Notably, glycemia

in D+SSTR2a was unaltered, which demonstrates that SSTR2 antagonist administration did

not affect glycemia during basal conditions.

Plasma glucagon levels remained unaltered at all time points in N and D groups during the

control experiments. However, Ctrl:D+SSTR2a demonstrated very modest, transient

increases in plasma glucagon at 40, 60, and 90 min (Table 4-4). AUC calculations revealed

no significant difference in overall glucagon in Ctrl:D and Ctrl:D+SSTR2a in the absence of

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hypoglycemia (P=0.12). Plasma corticosterone, epinephrine, and norepinephrine levels also

remained similar in all groups (Table 4-4). These controls demonstrate that SSTR2

antagonist, per se, did not elicit hyperglycemia or marked, sustained elevations of stress

hormones.

4.3.4 Pancreatic glucagon and pancreatic and plasma somatostatin measurements

Pancreatic glucagon protein content increased 50% in D compared to N in both

hypoglycemia and control studies (P<0.02) and was similar in both D and D+SSTR2a

(Figure 4-12A). Similarly, pancreatic somatostatin protein content following hypoglycemia

was increased by 65% in D rats as compared to N rats (P<0.02) and was not affected by

SSTR2 antagonist (Figure 4-12B). All three control groups showed no statistical difference

in pancreatic somatostatin content (Ctrl:N vs Ctrl:D: P=0.09; Ctrl:D vs Ctrl:D+SSTR2a:

P=0.07). Plasma somatostatin concentrations taken at euthanasia showed a similar trend

to pancreatic somatostatin: following hypoglycemia, plasma somatostatin increased 60%

increase in D, but this failed to reach statistical significance (P=0.08, Figure 4-12C).

4.3.5 Liver PEPCK1 and G6Pase mRNA expression

To determine whether the normalization of glucagon and corticosterone responses to

hypoglycemia in SSTR2 antagonist-treated diabetic rats also affected the expression of

these gluconeogenic enzymes, despite the lack of change in glucose infusion rates, real-time

PCR was used to quantitatively measure the levels of mRNA expression in the livers of the

high dose insulin group (10 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist). There was no

statistical difference in PEPCK1 expression between all groups, regardless of diabetes,

hypoglycemia, and SSTR2 antagonist administration (Table 4-5). In control groups, G6Pase

expression was 6-fold lower in Ctrl:D as compared to Ctrl:N (P<0.0001). Ctrl:D+SSTR2a

showed increased G6Pase expression as compared to Ctrl:D (P<0.05) during basal

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conditions. Insulin-induced hypoglycemia significantly suppressed G6Pase expression in N

(P<0.0001) and D+SST2Ra (P<0.007).

4.3.6 Effect of SSTR2 antagonist in non-diabetic rats during hypoglycemia

We assessed the effect of SSTR2 antagonist during hypoglycemia in non-diabetic rats.

Basal metabolic parameters of these rats (Table 4-6) were similar to non-diabetic controls.

Blood glucose levels during hypoglycemia clamp experiments were the same in N and

N+SSTR2a at each of the dosing combinations of insulin to induce hypoglycemia and SSTR2

antagonist tested (Figure 4-13A-C). At 1500 nmol/kg/h, SSTR2 antagonist had no effect on

glucagon responses to hypoglycemia in non-diabetic rats (Figure 4-14A). At 3000

nmol/kg/h, SSTR2 antagonist had an opposite effect and strikingly decreased plasma

glucagon levels in response to hypoglycemia at the 40, 60, 90, and 120 min time points

(Figure 4-14B, C). This suppressive effect of 3000 nmol/kg/h SSTR2 antagonist in non-

diabetic rats was more marked at the higher dose of insulin (10 U/kg, Figure 4-14B) than at

the lower dose of insulin (5 U/kg, Figure 4-14C). SSTR2 antagonism had no effect on

corticosterone responses to hypoglycemia in both of the doses tested (Figure 4-14D-F).

N+SSTR2a rats had similar pancreatic and plasma somatostatin levels as N following

hypoglycemia (Table 4-7).

4.4 Discussion

4.4.1 Role for SSTR2 in the enhancement of glucagon counterregulation

The attenuation or absence of the glucagon response to hypoglycemia in diabetes is well

established (165;166). We demonstrate for the first time that antagonism of SSTR2

enhances hypoglycemia-stimulated, but not basal, glucagon and corticosterone release in

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diabetic rats. In the present study, administration of SSTR2 antagonist (3000 nmol/kg/h)

fully restored glucagon counterregulation to hypoglycemia in diabetic rats induced with

either high (10 U/kg) or low (5 U/kg) dose insulin.

The inhibitory effect of somatostatin on -cell glucagon release via SSTR2 is well-

established in both rodents (389;531;532) and humans (385;533). In very early studies,

anti-somatostatin antibodies were shown to increase stimulated glucagon release from

isolated, non-diabetic islets (279), thus demonstrating that blocking the actions of

somatostatin could enhance glucagon secretion. A recent study demonstrated that global

somatostatin knockout increased nutrient-stimulated, but not basal, glucagon secretion

compared with wild-type mice in vivo and in isolated islets, the latter implicating the

profound role of locally released somatostatin on stimulated glucagon secretion (534). In

addition, in STZ-treated rats treated with insulin, it was demonstrated that the glucagon

response to insulin-induced hypoglycemia can be enhanced by pancreatic infusion and

subsequent switch-off (at hypoglycemia) of somatostatin, thereby supporting the notion

that glucagon counterregulation can be enhanced by deactivating intrapancreatic signals

suppressing -cells (179). The localization of particular receptor subtypes on different islet

cell types allows for specific receptor agonists/antagonist to exert specific effects on the

cells. In rodent islets, SSTR2 are found nearly exclusively on glucagon-secreting -cells

while SSTR5 are found on -cells (388;389). Studies in whole-body somatostatin receptor-

type-2 knockout (SSTR2 KO) mice showed that stimulated glucagon secretion was

approximately 2-fold greater in islets isolated from SSTR2KO mice than wild-type (389). In

humans, somatostatin also exerts its inhibitory effects on glucagon secretion via SSTR2 on

-cells (385;535). An in vitro study in human islets demonstrated that an agonist to the

SSTR2 was more potent than other receptor subtype agonists at inhibiting arginine-

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stimulated glucagon secretion and that a dose-dependent reversal of SSTR2-agonist-

induced glucagon suppression was achieved by using the same SSTR2 antagonist as the

present study (386). The findings from this study in human islets suggest that our present

work using this SSTR2 antagonist may also be relevant to human studies. Taken together,

these studies support the role of somatostatin acting via the SSTR2 in inhibiting stimulated

glucagon secretion.

We demonstrate that pancreatic content of glucagon remains elevated in diabetic rats even

subsequent to hypoglycemia. This supports our idea that during hypoglycemia, there is a

defect in the secretory mechanism of the diabetic -cell where glucagon remains present,

which contributes to impaired glucagon counterregulation. We speculate that this defect is

due, at least in part, to increased pancreatic somatostatin and increased insulin sensitivity

of diabetic -cells. In addition, we observe that pancreatic glucagon content is unchanged

with SSTR2 antagonist treatment, yet diabetic rats which received SSTR2 antagonist had

markedly enhanced glucagon release in response to hypoglycemia. This may suggest that

that the amount of glucagon secreted in response to hypoglycemia represents only a small

amount of glucagon stored in the pancreas or that glucagon is quickly re-synthesized after

its secretion in diabetic rats that received SSTR2 antagonist.

Presently, we observe that pancreatic somatostatin remains elevated in diabetic rats

following hypoglycemia as previously reported (174). It is because diabetic rats have

elevated somatostatin that we decided to double our initial dose of SSTR2 antagonist from

our pilot studies. Elevated pancreatic and circulating somatostatin has been reported in

various diabetic species, including type 1 diabetic humans, STZ-diabetic and alloxan-

diabetic animals, and NOD mice, but BB rats are one diabetic model in which pancreatic

somatostatin is not increased (536-539). Untreated diabetic BB rats demonstrated reduced

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concentration of pancreatic somatostatin (540). However, circulating somatostatin remains

elevated in BB rats (537;541). BB rats also have impaired glucagon and catecholamine

responses to hypoglycemia (263;542;543). Thus, increased pancreatic somatostatin is not

the only factor for impaired counterregulation. In the present study, there was a trend for

increased plasma somatostatin in diabetic groups, although this did not reach statistical

significance (Figure 4-12). Interestingly, however, diabetic rats in the control group under

basal conditions did not exhibit elevated somatostatin. It has been reported that fasting for

15-h (544) and 48-h (545) decreased pancreatic somatostatin levels in rats. This effect of

fasting may be biphasic since with prolonged fasting of 72 h in rats, pancreatic somatostatin

levels eventually become elevated (544). At the time of euthanasia when pancreata were

harvested, these rats would have been fasted for 24 h. Thus, the reduction in pancreatic

somatostatin under basal conditions could be due to a more profound effect of fasting on

diabetic groups. When we combined the data within each treatment group for all rats that

were subjected to hypoglycemia Table 4-7, we can observe a significant effect of diabetes in

elevating both pancreatic (P<0.0004) and plasma somatostatin (P<0.004) levels. Even if

somatostatin is not excessive, suppression of somatostatin may be useful to alleviate

further inhibition on diabetic -cells, which are already more sensitive to inhibition by

exogenous insulin during hypoglycemia.

Some may not have considered increased somatostatin to be a strong candidate for

impaired glucagon responsiveness because somatostatin-secreting -cells have been

reported to be downstream of glucagon-secreting -cells in the islet microcirculation of non-

diabetic rats (79;284). This differed from earlier views that suggested a parallel distribution

of -cells and -cells in all species and in which -- represented the core-to-periphery

arrangement of islet cells (276). Since pancreata from non-diabetic rats perfused with

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SSTR2 antagonist indeed show increased stimulated glucagon secretion (392), this supports

the role of somatostatin in the local regulation of glucagon release via the microcirculation

or interstitial interactions. In STZ-diabetic rats, however, this architecture of islet cell type

is altered such that -cells are also distributed in central portions of islet cells (546). The

arrangement of human endocrine islet cells is likewise more disperse throughout the islet

(304;547). Furthermore, immunofluorescent methods in pancreata of STZ-diabetic rats, as

well type 1 diabetic humans, revealed hyperplasia and hypertrophy of pancreatic -cells

(439). These findings suggest that paracrine actions of islet hormones may be altered in

diabetes such that -cell release of somatostatin may very well regulate glucagon secretion.

4.4.2 Potential role for SSTR2 in insulin secretion?

In some studies in human perfused pancreas (548) and of incubations of cloned hamster -

cell lines (549), it was demonstrated that activation of SSTR2 could inhibit insulin secretion,

which may lead to speculation that SSTR2 antagonism may promote insulin release. The

laboratory of Efendic also demonstrated a moderate (~30%) increase in arginine-stimulated

insulin secretion with the same SSTR2 antagonist (PRL-2903) in batch incubated islets of

non-diabetic rats (392). It should be noted, however, that the increase in glucagon

secretion (5-fold) under the same conditions was much greater thus demonstrating a much

larger effect of the SSTR2 antagonist on stimulated glucagon, rather than insulin, secretion.

There was no effect of the SSTR2 antagonist in insulin secretion in perifused isolated islets

or perfused pancreas. Although this SSTR2 antagonist has a 20-fold greater binding affinity

for SSTR2 than SSTR5 (expressed on rodent -cells), high concentrations of SSTR2

antagonist may exert a non-specific effect if sufficient amounts of a native competitor (i.e.

endogenous somatostatin) are not present. This concept of non-specific role of SSTR2 in

promoting insulin secretion will be discussed further in section 4.4.9 in the interpretation of

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our data for SSTR2 antagonism in non-diabetic rats during hypoglycemia. In contrast, our

experiments conducted under basal conditions do not demonstrate an effect on insulin

release with SSTR2 antagonism, presumably primarily because of STZ-diabetic rats have

markedly less pancreatic insulin due to -cell destruction, and also because SSTR5 rather

than SSTR2 is the predominant isoform of SSTR on rodent -cells (387). It is possible that

SSTR2 antagonism may have some effect on insulin secretion in humans since some SSTR2

are detected human, but not rodent, -cells, (385). This expression is relatively low as it

was reported in pancreatic islets of non-diabetic humans that approximately 45% of -cells

expressed SSTR2 whereas approximately 100% and 90% expressed SSTR1 and SSTR5,

respectively. Furthermore, SSTR2 antagonism is unlikely to exacerbate hypoglycemia if

used clinically in diabetic humans since individuals with type 1 diabetes will not have

functioning insulin-secreting -cells.

4.4.3 Role for SSTR2 in the enhancement of corticosterone counterregulation

Whereas glucagon is important during the early fall in plasma glucose, catecholamines and

glucocorticoids become increasingly important at lower glucose levels and during prolonged

hypoglycemia (3;232). Dysregulation of the hypothalamo-pituitary-adrenal (HPA) axis can

contribute to defective counterregulation, since it not only regulates glucocorticoid secretion

but also affects epinephrine secretion (202;220;550-554). Glucocorticoids increase liver

sensitivity to glucagon and epinephrine as well as decrease hepatic and peripheral insulin

sensitivity (555;556). Presently, we demonstrate that peripheral administration of SSTR2

antagonist normalizes the attenuated corticosterone response to hypoglycemia in diabetic

rats, which may be due to a central effect of somatostatin antagonism or to a peripheral

effect of SSTR2 antagonism at the level of the pituitary or adrenal glands. The attenuated

corticosterone response in diabetic rats occurred in both high (10 U/kg) and low (5 U/kg)

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dose insulin-induced hypoglycemia experiments in which basal corticosterone was either

elevated or similar to non-diabetic rats. The markedly blunted corticosterone responses of

diabetic rats as compared with non-diabetic supports previous findings (202;205). In our

low-dose insulin-induced hypoglycemia experiments, plasma insulin levels were similar at all

time after insulin administration. Despite the fact that all rats in the high dose insulin

experiments received 10 U/kg, plasma insulin levels at t=60 and 180 min were

approximately 50% elevated in non-diabetic as compared to diabetic controls. This may be

due to enhanced clearance of exogenous insulin by diabetic rats. It has been reported that

insulin dose and consequent plasma levels, per se, affect hormonal responses to

hypoglycemia. In non-diabetic humans, a 6-fold higher insulin level significantly increased

cortisol and catecholamine, but not glucagon, responses by 58% and 33%, respectively, to

hypoglycemia despite an equivalent level of hypoglycemia, but this amplifying effect of

insulin was lost in subjects with type 1 diabetes (209;210). Thus, the higher plasma insulin

levels in our non-diabetic rats of the high-dose insulin-induced hypoglycemia study may be

of concern. In our studies, high-dose insulin yielded a 2-fold greater peak plasma insulin

level as compared to low-dose insulin experiments in our non-diabetic rats; however, peak

cortisol, epinephrine, and norepinephrine levels were similar. Glucagon responses were

slightly, but not significantly, greater in high-dose insulin as compared with low-dose insulin

regimes. Thus, in our studies, it is unlikely that insulin, per se, caused an amplifying effect

of counterregulatory hormones to hypoglycemia, and thus the difference in peak plasma

insulin levels between non-diabetic and diabetic controls in the high-dose insulin

experiments would not account for the observation of attenuated corticosterone (or

glucagon) counterregulation in diabetic rats.

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Basal corticosterone levels are often reported to be elevated in diabetic rats, an effect which

is attributed to increased HPA activity due to molecular HPA dysregulation (202;205;557).

Increased basal corticosterone in diabetes was observed in our high dose insulin

hypoglycemia experiments, but not in our low dose insulin hypoglycemia or control

experiments. We attribute the lack of elevated corticosterone to increased basal

corticosterone levels of our non-diabetic controls since these values were augmented

approximately 4-fold relative to our non-diabetic rats in the high dose insulin group, which

had a comparatively larger sample size. This increase in basal corticosterone in non-

diabetic rats may have been due to unwanted stress effects, but since basal epinephrine

levels remained low, any unwanted stress was likely very mild. A circadian effect is also

possible if basal hormone samples of thee non-diabetics were obtained slightly later in the

day. It is known that in the non-diabetic rat, a nocturnal animal, the corticosterone nadir

occurs in the morning and peaks in the evening (558). However, diabetic rats (559), as

well as humans (560), disrupted circadian patterns of corticosterone/cortisol secretion, with

elevated levels during the trough and normal or slightly elevated values during peak

secretion. However, this is unlikely to have affected our results since the patterns of

corticosterone responses to hypoglycemia were similar in both sets of hypoglycemia

experiments.

The role of somatostatin on the regulation of corticosterone is less well-documented than its

role on suppressing glucagon release. However, somatostatin also inhibits the secretion of

both CRH and ACTH (271;294), which regulate corticosterone secretion; thus, somatostatin

inhibits the function of the HPA axis. SSTR2 have been localized in several brain regions

and specifically in areas pertaining to HPA activity – the hypothalamic paraventricular

nucleus (420) and ACTH cells in the anterior pituitary (561). It is unknown whether this

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antagonist crosses the blood-brain-barrier. However, a cyclic octapeptide with a similar

chemical structure, CTAP, is known to enter the brain when administered peripherally (562).

The corticosterone response to hypoglycemia is often impaired in diabetes

(202;205;206;225;563). Our laboratory has previously shown that carbachol injected

intracerebroventricularly into the third ventricle (a model of stress) increases the release of

cortisol in non-diabetic and diabetic dogs (169). However, when somatostatin was given

acutely with carbachol, cortisol responses were abolished in both normal and diabetic dogs

(528). Furthermore, somatostatin has been demonstrated to inhibit arginine vasopressin

release in response to hypoglycemia (564). Arginine vasopressin is one of the

neuropeptides involved in the activation of the HPA axis during stress. This supports the

idea that the SSTR2 antagonist could improve counterregulation through a central pathway

and could be a mechanism whereby the SSTR2 antagonist also markedly increases the

corticosterone response to hypoglycemia in diabetic rats. Along with other subtypes, SSTR2

are normally expressed in the pituitary (561). Pituitaries of SSTR2 knockout mice release

more ACTH in vitro (565), and corticosterone levels are elevated in SST knockout mice

(566), which supports the role of endogenous somatostatin on regulating corticosterone

release. An alternative possibility is an enhancement of corticosterone through

somatostatin receptors in the adrenal cortex. In the normal adrenal gland, all five subtypes

of SSTR are identified although SSTR2 and SSTR4 are most highly expressed (567;568).

However, despite consistent detection of SSTR2 in the zona glomerulosa (390;397-399), its

detection is not as consistent in the corticosterone-synthesizing fasciculata and reticularis

zonae of the adrenal cortex (390;569).

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4.4.4 Epinephrine responses to hypoglycemia were unaffected

In the present study, we observed no difference in the epinephrine response to

hypoglycemia in non-diabetic and three-week STZ-diabetic rats. Although this lack of an

epinephrine defect contrasted with our prior experiments in which intravenous insulin

infusion was used to induce hypoglycemia (202;205), it is consistent with other

observations in which hypoglycemia was induced with a subcutaneous insulin injection

(174), suggesting that the route of insulin administration may also play a role in the

epinephrine response. An effect of the SSTR2 antagonist on epinephrine levels in diabetic

rats was not observed, either because somatostatin has no role in epinephrine

counterregulation to hypoglycemia or because the maximal physiological epinephrine

response was already achieved. It would be of future interest to evaluate the effect of

SSTR2 antagonism in a diabetic model in which a defect of epinephrine is observable (e.g.

diabetes of longer duration (570)) to determine whether somatostatin may also play a role

in hypoglycemia-stimulated epinephrine release.

Somatostatin inhibits glucagon secretion in response to various stimuli, and it also blocks

the effect of epinephrine to stimulate glucagon release as was demonstrated by the

exogenous administration of somatostatin to isolated perfused rat pancreas (571).

Somatostatin and epinephrine share common mechanistic pathways by which they regulate

glucagon release from the -cell. Somatostatin activation of SSTR2 inhibits adenylate

cyclase activity, thus reducing cAMP levels and subsequently inhibits protein kinase A (PKA)-

stimulated glucagon release (572-574). PKA is a serine/threonine kinase that

phosphorylates substrate proteins at serine or threonine sites (575). In contrast,

epinephrine activation of 1- and -adrenergic receptors on -cells stimulates adenylate

cyclase activity, in turn increasing cAMP levels and promoting PKA-stimulated glucagon

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release (576-578). Somatostatin abolishes L-type Ca2+ channel-dependent release of

glucagon (579), whereas epinephrine enhances Ca2+ influx through L-type Ca2+ channels

which increased intracellular Ca2+ levels to promote glucagon release (577). Somatostatin

activates calcineurin, a serine/threonine phosphatase that “deprimes” glucagon-containing

granules and thus inhibits glucagon release (579), whereas epinephrine accelerates the

mobilization of glucagon-containing granules, increasing the number of readily releasable

granules by 5-fold (577). Thus, epinephrine can help to enhance glucagon secretion during

hypoglycemia, but excessive somatostatin may hinder this enhancement. In our present

studies, a robust glucagon response to hypoglycemia in our non-diabetic rats was

accompanied with a robust epinephrine response. In our diabetic rats, however, the

glucagon response to hypoglycemia was markedly abolished despite epinephrine responses

of the same magnitude as non-diabetic rats. When diabetic rats were treated with SSTR2

antagonist, glucagon responses to hypoglycemia were normalized. Thus, we presently

hypothesize that SSTR2 antagonism may also alleviate the suppressive effects of

somatostatin on the potentiating effects of epinephrine on glucagon release in response to

hypoglycemia.

4.4.5 SSTR2 antagonist demonstrated no effect on growth hormone

Presently, we see no effect of SSTR2 antagonism on growth hormone release during the 1-h

basal period or during 3-h hypoglycemia when SSTR2 antagonist was given at high dose.

Furthermore, no increase of growth hormone is observed in response to hypoglycemia.

Since growth hormone is known to play a role in prolonged hypoglycemia (143;580), the

present hypoglycemia experiments may not have been of sufficient duration to stimulate a

growth hormone response. Another possibility for the lack growth hormone increase is that

plasma epinephrine levels were markedly increased during hypoglycemia. This elevation of

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epinephrine could have suppressed growth hormone, as could an epinephrine-induced

increase in free fatty acid levels, both of which are known to suppress growth hormone

secretion (581). Also, it should be noted that samples were obtained at very few time

points (four time points over a four hour experiment). Since growth hormone release is

pulsatile (581), frequent blood sampling is required to accurately measure growth hormone

response. Thus, the growth hormone data obtained in the present study may not truly be

indicative of the pattern anticipated in response to hypoglycemia. In response to prolonged

hypoglycemia, growth hormone counterregulation can contribute to the promotion of

glycemic recovery by suppressing insulin-mediated glucose utilization and increasing

glucose release into the circulation (582;583). When growth hormone secretion is inhibited,

liver and peripheral insulin sensitivity decreases (500;584), and nonesterified fatty acid

levels increase (500). SSTR5 is the predominant isoform of SSTR expressed in rat pituitary,

followed by the expression of SSTR2. Immunohistochemical colocalization studies

demonstrated that SSTR5 is expressed in 86% of rat GH-positive cells as compared to

SSTR2 in 42% of rat somatotrophes (379). Furthermore, it has demonstrated that for

somatostatin to effectively inhibit growth hormone secretion, the SSTR5 must also be

involved (431), and heterodimerization and activation of both SSTR2 and SSTR5 results in

synergistic GH suppression (432;585). Thus, it is not surprising that our selective SSTR2

antagonist, which does not have a high affinity for SSTR5, does not affect growth hormone

release in our experiments. However, it should be noted that growth hormone is very

pulsatile in its release and that we took measurements at relatively few time points and

may have missed periods of plasma differences in growth hormone secretion.

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4.4.6 SSTR2 antagonism does not cause hyperglycemia or undesired stress

We also demonstrate that SSTR2 antagonist does not affect glycemia or markedly alter

counterregulatory hormones in the absence of hypoglycemia. The control experiments in

the absence of insulin were performed to examine the effects of SSTR2 antagonist, per se,

during basal conditions.

Somatostatin analogues were clinically used to help lower blood glucose levels in diabetic

patients (586;587). Also, the anti-diabetic activity of a SSTR2 agonist was demonstrated in

rodent models of type 2 diabetes in which the agonist lowered plasma glucagon and glucose

levels (588). Thus, it was important to evaluate the effect of the SSTR2 antagonist when

administered during the basal state if SSTR2 antagonism could have any putative clinical

role as a therapy for hypoglycemia in the diabetic population. Since basal

hyperglucagonemia can persist despite elevated plasma somatostatin levels in diabetes, the

role of somatostatin-induced glucagon inhibition has been questioned (489). Presumably,

some could speculate that the SSTR2 antagonist may promote basal hyperglycemia, which

is undesired in diabetic patients. However, this can be reconciled with understanding an

action of somatostatin receptor-coupled signalling in the -cell: somatostatin via SSTR2

inhibits Ca2+-induced glucagon exocytosis via Gi2-dependent activation of calcineurin, a

serine/threonine protein phosphatase which “deprimes” glucagon-containing granules

associated with L-type Ca2+ channels but does not affect the granules associated with N-

type Ca2+ channels, which mediate basal glucagon secretion (577;579;589). In addition to

this mechanism, somatostatin can also inhibit adenylate cyclase activity, consequently

reducing cAMP levels and protein kinase A (PKA)-stimulated glucagon secretion, as well as

activate G-protein-coupled K+ channels on -cells, which causes hyperpolarization of the

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plasma membrane and subsequent inhibition of -cell electrical activity and glucagon

release (589).

Presently, we demonstrate that antagonism of SSTR2 enhances hypoglycemia-stimulated,

but not basal, glucagon release in diabetic rats. In vitro experiments in pancreatic islets of

non-diabetic mice demonstrate that blocking somatostatin receptor type 2 using SSTR2

antagonist stimulated glucagon secretion in low glucose conditions but did not affect

glucagon secretion at “normal” glucose conditions of 7 mM (590). Administration of SSTR2

antagonist in perfused non-diabetic rat pancreas did not affect the inhibitory effect of

hyperglycemia on glucagon secretion (591), thus confirming that SSTR2 did not elicit

hyperglucagonemia when glucose concentrations were already high. Presently, we

demonstrate that SSTR2 antagonism during basal conditions in vivo yielded very small

transient increases of glucagon which did not cause hyperglycemia and did not alter

corticosterone or catecholamine levels, suggesting that this antagonist does not elicit

marked, non-stimulated secretion of counterregulatory hormones. However, it remains to

be investigated whether prolonged administration of SSTR2 antagonists will lead to

hyperglycemia. Studies in SSTR2 knockout mice showed no differences in fasting and fed

blood glucose levels, fasting and fed plasma glucagon levels, or responses to glucose

tolerance tests compared wild-type controls, but diet-induced obesity in these knockouts

caused fasting and fed hyperglycemia, fed hyperglucagonemia, and impaired glucose

tolerance compared to wild-types given the same high-fat diet (592). It remains to be seen

if a prolonged administration of SSTR2 antagonist in diabetic animals would exacerbate

diabetes.

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4.4.7 Glucose requirement during hypoglycemia

Glucose infusion rates and total glucose infused were decreased with SSTR2 antagonism

during low-dose insulin-induced hypoglycemia in the present study. A lower glucose

requirement during hypoglycemia is consistent with normalized glucagon and corticosterone

responses in the SSTR2 antagonist-treated diabetic group.

At the high insulin dose, there were no differences in glucose infusions rates between non-

diabetic and diabetic rats. With three weeks of chronic hyperglycemia in our STZ-diabetic

rats, it would be anticipated that some insulin resistance would develop, as has been

previously reported (205). However, glucose infusion rates were not lower in untreated

diabetic rats during hypoglycemia presumably because the expected insulin resistance of

diabetic rats was offset by the generally lower counterregulatory response. Furthermore,

the hypothesized reduction in glucose requirement in SSTR2 antagonist treated animals

may have been masked by the overwhelming maximal inhibitory effect of insulin to

suppress endogenous glucose production and increase glucose utilization. Our method of

inducing hypoglycemia with insulin injection creates a major perturbation of steady state

which renders classic glucose turnover techniques to measure glucose production and

utilization less reliable, especially during the rapid change of glycemia at the onset of insulin

injection in the present experiments, thus we used the exogenous glucose requirement and

expression of gluconeogenic enzymes as indicators of effects of improved counterregulation

on glucose homeostasis.

Despite using half the dose of insulin in our low-dose versus high-dose insulin experiments,

glucose infusion rates seemingly appear greater in groups induced with 5 U/kg insulin. We

attribute this to the fact that hypoglycemia was reached at an earlier time point in the low-

dose study (i.e. non-diabetic rats: at t=20 min for low-dose as compared to t=50 min for

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high dose insulin-induced hypoglycemia experiments; both diabetic groups: at t=25 min for

low-dose as compared to t=70 min for high-dose insulin-induced hypoglycemia

experiments), and thus the glucose requirement would be greater to maintain clamped

hypoglycemia for a longer duration. Alternatively, diabetic rats in the low-dose insulin

experiments may have had a greater exogenous glucose requirement since they had a lower

initial basal glycemia and may have been less insulin resistant than diabetic rats in the high-

dose group. Regardless, diabetic groups within each experiment were similar, and thus the

effects of SSTR2 antagonist treatment could be accurately compared.

4.4.8 Expression of gluconeogenic enzymes

Gluconeogenesis plays a role in restoring blood glucose levels in response to hypoglycemia

and is particularly relevant after an overnight fast in which glycogen stores are depleted in

rodents, resulting in gluconeogenesis rather than glycogenolysis as the main contributor to

glucose synthesis (593-595). Glucagon (596-598) and corticosterone (599-601) are well

known to induce the expression of PEPCK and G6Pase. Glucagon activates protein kinase A,

increasing cAMP levels, which induces gluconeogenic enzymes (596-598). Corticosterone

induces PEPCK (599) and G6Pase (600;601) expression by binding with glucocorticoid

receptors and activating glucocorticoid-responsive elements on the promoter regions of

these genes. Furthermore, the nuclear coactivator protein peroxisome proliferator activated

receptor- coactivator 1 (PGC-1) is induced by glucagon and corticosterone and promotes

increased transcription of both PEPCK and G6Pase by its interaction with HNF-4, thus

contributing as a key mechanism for controlling glucose production in hepatocytes (602).

Since glucagon and corticosterone responses to hypoglycemia were normalized in diabetic

rats with SSTR2 antagonism without an effect on glucose infusion rates at the high insulin

dose, we wished to determine if gene expression of these gluconeogenic enzymes was

110

increased and this would have served as better markers than glucose infusion rates of

counterregulatory hormone actions.

Hypoglycemia markedly decreased G6Pase mRNA expression in both non-diabetic and

diabetic rats, which is attributed to the large amount of insulin administered (up 200-fold

increase in plasma levels), which has a large suppressive effect on gluconeogenic enzyme

synthesis (596;603) and can override the stimulatory effect of increased counterregulatory

hormones (604). PEPCK1, the cytosolic form, is the predominant form in rodent liver (605).

Unlike G6Pase, expression of PEPCK1 was not decreased by hypoglycemia in the present

study. This is in accordance with another study in which epinephrine could overcome the

inhibitory effect of insulin on stimulating PEPCK1 expression in rat hepatocytes (606). Thus,

despite robust increases of glucagon and corticosterone during hypoglycemia, the

expression of PEPCK1 and G6Pase was not affected, perhaps masked by the stimulatory

effect of epinephrine or overwhelming inhibitory effect of insulin, respectively.

In the absence of hypoglycemia, G6Pase in diabetic rats was actually decreased compared

with non-diabetic rats, and SSTR2 antagonist treatment in diabetic rats partially increased

this expression. The modest, transient increase of plasma glucagon levels in the SSTR2

antagonist-treated group may be responsible for the moderate increase in G6Pase

expression in the present study. Modest increases in circulating glucagon are sufficient in

increasing glucose production in the short-term (i.e. 2 h) in partially insulin-deprived

humans (607), although in our study, hyperglycemia was not observed during the 4-h time-

frame of our experiments. Our basal gluconeogenic enzyme results are in contrast to the

popular view that expression of gluconeogenic enzymes PEPCK (608-610) and G6Pase

(601;611-614) are elevated with diabetic rodents, but are in line with a recent finding in

mildly hyperglycemic, overnight fasted, STZ-diabetic rats, (615). A shorter duration of

111

fasting (i.e. 6-h) in STZ-diabetic rats likewise yielded no differences in PEPCK1 and GP6Pase

expression despite elevated fasting plasma glucose levels and rates of endogenous glucose

production (615). In addition to suggesting the role of other key enzymes which may be

responsible for regulating gluconeogenesis, the authors suggest that increased PEPCK and

G6Pase expression are often associated with diabetic models with increased corticosterone

levels and demonstrate that expression does not increase from controls in STZ-diabetic rats

without hypercorticosteronemia. In our non-diabetic and diabetic rats which had similar

plasma corticosterone levels for the duration of the control experiments, we likewise do not

observe increased expression of gluconeogenic enzymes. Another possibility is that basal

gluconeogenic enzyme transcription is not increased in our diabetic rats despite lower

endogenous insulin because of hyperglycemia. At present, there is no known direct effect of

SSTR2 antagonism on hepatic gluconeogenesis. SSTR1 (271) and SSTR3 (373), but not

SSTR2, have been detected on hepatocytes, and the SSTR2 antagonist has no reactivity

with SSTR1 and 10-fold less binding affinity with SSTR3 (509). In isolated rat hepatocytes,

somatostatin has been reported to have no direct effects on basal glucose production

(616;617) on glucagon-stimulated glucose production, or on PEPCK and G6Pase activity

(617). Similarly, studies in conscious dogs have demonstrated that somatostatin does not

alter basal glucose production rates when the levels of insulin and glucagon are maintained

(618). Besides its use in pancreatic clamp studies to measure glucose turnover, the direct

effects of somatostatin effect on the liver are not well-documented aside from its use in liver

tumour cells or cirrhotic liver (418). Somatostatin bound to its receptors elicit their cellular

responses through G-protein coupling, including inhibition of adenylyl cyclase via inhibitory

G proteins (271), thus it is possible that somatostatin can lower cAMP levels directly within

hepatocytes, consequently blunting expression of gluconeogenic enzymes such as PEPCK

and G6Pase and thus affecting gluconeogenesis and subsequent hepatic glucose production.

112

However, there is no evidence that such a mechanism exists at the present time. However,

the modest, transient increase of plasma glucagon levels in the SSTR2 antagonist-treated

group may be responsible for the moderate increase in G6Pase expression observed in the

present study.

4.4.9 Effect of SSTR2 antagonism during hypoglycemia in non-diabetic rats

The main objective of this study was to evaluate the effect of SSTR2 blockade on hormone

counterregulation to hypoglycemia in diabetic rats. We postulated that elevated

somatostatin in diabetes contributes to this impairment, and SSTR2 antagonism in non-

diabetic rats provides a control during hypoglycemia so that the magnitude of the

impairment in diabetes that might be attributable to elevated somatostatin may be

assessed. Both groups of non-diabetic rats in this study have similar pancreatic and plasma

somatostatin levels (Table 4-7). We presently provide evidence that SSTR2 antagonist

treatment in non-diabetic rats does not increase glucagon or corticosterone responses to

hypoglycemia. This is in contrast to diabetic rats with elevated somatostatin in which

disinhibition of somatostatin acting via SSTR2 normalized the defects of both glucagon and

corticosterone counterregulation. We assessed the effects of diabetes and administration of

SSTR2 antagonist on pancreatic and plasma somatostatin levels by combining available data

of the groups form the hypoglycemia experiments (Table 4-7). Using Two-Way ANOVA

statistical analysis, it can be shown that there is an effect of diabetes, but not SSTR2

antagonist (P=0.51 for both), on both pancreatic somatostatin (P<0.0004) and plasma

somatostatin (P<0.004) values. Thus, data from the non-diabetic control for SSTR2

antagonism during hypoglycemia provides support for a role of elevated somatostatin in

diminishing glucagon and corticosterone responses to hypoglycemia in diabetic rats, but not

non-diabetic rats.

113

At 1500 nmol/kg/h, SSTR2 antagonist had no effect on glucagon responses to hypoglycemia

in non-diabetic rats. This supports the hypothesis that somatostatin does not play a role in

suppressing glucagon release in non-diabetic rats, but taken together with the data in

diabetic rats in which somatostatin is elevated, it demonstrates that disinhibition of the -

cell (which is has increased sensitivity to the suppressive effects of exogenous insulin)

during hypoglycemia enhances glucagon release in diabetes. However, it was very

surprising to observe that when a two-fold larger dose of SSTR2 antagonist was used in

non-diabetic rats, an effect opposite to that which would be expected of somatostatin

blockade ensued: glucagon responses were markedly attenuated rather than enhanced.

This effect could have been mediated by a stimulated release of insulin resulting from a

non-specific effect of a high dose SSTR2 antagonist on -cells (which express SSTR5 and

not SSTR2 in rats), which could locally act to suppress glucagon release. It is possible that

when high doses of this antagonist are used in the absence of a sufficient amount of a

competitor (i.e. non-diabetic rats have significantly less somatostatin than diabetic rats),

binding to the SSTR5 on -cells may occur, which could disinhibit the -cell from the

suppressive effects of somatostatin and promote insulin secretion. Cejvan et al. reported

that this SSTR2 antagonist under certain conditions at high dose modestly increased insulin

secretion in non-diabetic rat islets when it should have had no effect (392). A similar

concept has previously been suggested by Gromada et al. (589) who used a high dose of

SSTR2 antagonist in studies of isolated rat -cells. In this in vitro setting in which the

purified -cell fraction did not contain any detectable amounts of endogenous somatostatin,

the SSTR2 antagonist applied at a comparatively high dose (2 M) increased insulin

secretion, which led to the authors cautioning against the use of these compounds in the

absence of native competitive receptor agonists (589). Thus, despite the fact that our

SSTR2 antagonist binds with 20-fold lower affinity to SSTR5 than SSTR2, thus suppressing

114

glucagon release, it is possible that -cell release of insulin may have been stimulated by

high dose SSTR2 antagonist in non-diabetic rats of our study. Non-diabetic rats did not

have augmented pancreatic somatostatin as diabetic rats, and the excessive amount of the

peptide may have suppressed the -cell rather than disinhibiting it. Future analysis using a

C-peptide assay could determine whether SSTR2 antagonist stimulates insulin release in

non-diabetic rats. This would not occur in diabetic rats, which have both elevated

somatostatin and markedly reduced -cell function. Alternatively, it is also possible that at

high doses, our peptide antagonist could have partial agonist activity. One group has

cautioned that certain competitive peptide analogues could exert opposite effects when used

at higher concentrations (619).

4.5 Study 1 Conclusions

In conclusion, we demonstrate that glucagon and corticosterone counterregulatory hormone

responses to hypoglycemia can be normalized in diabetic rats with specific antagonism of

the SSTR2, which does not elicit hyperglycemia or substantial elevations in

counterregulatory hormones during basal conditions. Our results therefore suggest an

important role for increased pancreatic, and possibly circulating, somatostatin in defective

glucagon and corticosterone counterregulation in diabetes but do not rule out other

mechanisms of impairment. Under conditions of lower dose insulin injection, SSTR2

antagonism reduced the glucose requirement during hypoglycemic clamp, which is

indicative of a lower risk of hypoglycemia. In this model of diabetes, epinephrine and

norepinephrine counterregulation to hypoglycemia induced with both high and low doses of

insulin were similar in both non-diabetic and diabetic rats. Growth hormone levels were not

affected by SSTR2 administration.

115

Figure 4-1. Study 1 protocol design.

116

Figure 4-2. Schematic representation of A) hypoglycemia clamp experiments (solid lines) and B) control experiments during basal conditions in the absence of exogenously administered insulin (dashed lines). In (A), “x” represents dose of insulin (high dose = 10 U/kg, low dose = 5 U/kg) and “y” represents dose of SSTR2 antagonist (pilot study = 1500 nmol/kg/h, else = 3000 nmol/kg/h). In (B), no insulin or glucose infusions are administered in these experiments under basal conditions. White diamonds (◊): N; black squares (): D; grey circles (): D+SSTR2a.

117

Figure 4-3. Pilot study: Plasma hormone levels during hypoglycemia clamp (10 U/kg insulin, 1500 nmol/kg/h SSTR2 antagonist). A) Glucagon. B) Corticosterone. White diamonds (◊): N; black squares (): D; grey circles (): D+SSTR2a. White bars: N; black bars: D, grey bars: D+SSTR2a. Data are represented as means ± SEM. *P<0.05 D vs N. †P<0.05 D vs D+SSTR2a.

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Figure 4-4. Blood glucose levels during hypoglycemia clamp (10 U/kg insulin, 3000 nmol/kg SSTR2 antagonist). White diamonds (◊): N; black squares (): D; grey circles (): D+SSTR2a. Data are represented as means ± SEM.

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Figure 4-5. Plasma hormone levels during hypoglycemia clamp (10 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist). A) Glucagon. B) Corticosterone. C) Glucose infusion. White diamonds (◊): N; black squares (): D; grey circles (): D+SSTR2a. White bars: N; black bars: D, grey bars: D+SSTR2a. Data are represented as means ± SEM. *P<0.002 D vs N. †P<0.05 D vs D+SSTR2a.

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Figure 4-6. Plasma A) epinephrine and B) norepinephrine levels during hypoglycemia clamp (10 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist). White diamonds (◊): N; black squares (): D; grey circles (): D+SSTR2a. Data are represented as means ± SEM.

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Figure 4-7. Blood glucose levels during hypoglycemia clamp using low insulin dose (5 U/kg insulin, 3000 nmol/kg SSTR2 antagonist). White diamonds (◊): N; black squares (): D; grey circles (): D+SSTR2a. Data are represented as means ± SEM.

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Figure 4-8. Plasma hormone levels during hypoglycemia clamp (5 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist). A) Glucagon. B) Corticosterone. C) Glucose infusion. White diamonds (◊): N; black squares (): D; grey circles (): D+SSTR2a. White bars: N; black bars: D, grey bars: D+SSTR2a. Data are represented as means ± SEM. *P<0.05 D vs N. †P<0.05 D vs D+SSTR2a. A. B.

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Figure 4-9. Plasma A) epinephrine and B) norepinephrine levels during hypoglycemia clamp (5 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist). White diamonds (◊): N; black squares (): D; grey circles (): D+SSTR2a. Data are represented as means ± SEM. A. B.

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Figure 4-10. Plasma growth hormone levels during hypoglycemia clamp (5 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist). White diamonds (◊): N; black squares (): D; grey circles (): D+SSTR2a. Data are represented as means ± SEM.

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Figure 4-11. Blood glucose levels during control experiments in the absence of insulin (0 U/kg, 3000 nmol/kg/h). White diamonds (◊): N; black squares (): D; grey circles (): D+SSTR2a. Data are represented as means ± SEM.

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Figure 4-12. Pancreatic protein content of A) glucagon and B) somatostatin, and C) plasma somatostatin, following hypoglycemia clamp (10 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist) and control experiments in the absence of insulin (0 U/kg, 3000 nmol/kg/h). Solid bars represent hypoglycemia experiments. Hashed bars represent control experiments. White bars: N; black bars: D, grey bars: D+SSTR2a. Data are represented as means ± SEM. *P<0.02 D vs N. A. B. C.

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Figure 4-13. Blood glucose levels during hypoglycemia clamp in non-diabetic rats at the various dosing combinations of insulin and SSTR2 antagonist. A) 10 U/kg insulin, 1500 nmol/kg/h SSTR2 antagonist; B) 10 U/kg insulin 3000 nmol/kg/h SSTR2 antagonist; (C) 5 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist. White diamonds (◊): N; grey triangles (): N+SSTR2a. Data are represented as means ± SEM.

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Figure 4-14. Effect of SSTR2 antagonist on plasma glucagon A-C) and corticosterone D-F) at the various dosing combinations of insulin and SSTR2 antagonist in non-diabetic rats. TOP (A, D): 10 U/kg insulin, 1500 nmol/kg/h SSTR2 antagonist; MIDDLE (B, E): 10 U/kg insulin 3000 nmol/kg/h SSTR2 antagonist; BOTTOM (C, F): 5 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist. White diamonds (◊): N; grey triangles (): N+SSTR2a. Data are represented as means ± SEM. *P<0.05 N vs N+SSTR2a. †P<0.005 N vs. N+SSTR2a.

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Table 4-1. Metabolic parameters of rats that underwent hypoglycemia clamps or control experiments in the absence of insulin. Fed blood glucose, body weight, and food intake were measured daily, and the table below shows the mean of values averaged over the last week before the experimental day. Daily blood glucose measurements were obtained using a glucometer. Data are represented as means ± SEM. *P<0.0002 D vs N in the respective experiment. **P<0.03 D vs N in the respective experiment. †P<0.04 D vs D+SSTR2a in the respective experiment.

fed blood glucose (mM)

body weight (g)

food intake (g)

(pilot study) hypoglycemia clamps (10 U/kg; 1500 nmol/kg/h)

N (n=5)

5.3 ± 0.2 408 ± 5 18 ± 2

D (n=3)

28.1 ± 1.8 * 340 ± 7 * 42 ± 3 *

D+SSTR2a (n=3)

26.3 ± 2.7 351 ± 9 36 ± 3

hypoglycemia clamps (10 U/kg; 3000 nmol/kg/h)

N (n=14)

5.2 ± 0.1 435 ± 7 23 ± 1

D (n=14)

22.4 ± 1.2 * 365 ± 5 * 37 ± 2 *

D+SSTR2a (n=18)

22.8 ± 1.1 355 ± 6 30 ± 1 †

hypoglycemia clamps (5 U/kg; 3000 nmol/kg/h)

N (n=5)

5.6 ± 0.3 400 ± 15 19 ± 3

D (n=4)

24.8 ± 3.5 * 323 ± 20 ** 32 ± 4 **

D+SSTR2a (n=4)

25.6 ± 1.0 331 ± 25 35 ± 3

controls in absence of insulin (0 U/kg; 3000 nmol/kg/h)

Ctrl:N (n=7)

5.8 ± 0.1 419 ± 6 18 ± 1

Ctrl:D (n=7)

24.7 ± 1.3 * 378 ± 9 ** 36 ± 3 **

Ctrl:D+SSTR2a (n=7)

26.4 ± 1.9 368 ± 15 38 ± 3

130

Table 4-2. Basal plasma counterregulatory hormone levels for rats that underwent hypoglycemia clamp (10 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist) and control experiments in the absence of insulin (0 U/kg, 3000 nmol/kg/h) at time = -60 min. “N/A” denotes time points at which samples for the particular hormone were not obtained. Data are represented as means ± SEM. *P<0.01 D vs N.

Basal plasma hormone concentration

glucagon (pg/mL)

corticosterone (ng/mL)

epinephrine (pg/mL)

norepinephrine (pg/mL)

(pilot study) hypoglycemia clamps (10 U/kg insulin, 1500 nmol/kg/h SSTR2 antagonist)

N 31 ± 9 134 ± 34 N/A N/A

D 71 ± 14 311 ± 32 N/A N/A

D+SSTR2a 46 ± 2 309 ± 133 N/A N/A

hypoglycemia clamps (10 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist)

N 32 ± 4 42 ± 9 55 ± 11 162 ± 26

D 51 ± 5 * 156 ± 26 * 148 ± 28 * 211 ± 35

D+SSTR2a 65 ± 5 148 ± 33 134 ± 57 334 ± 60

hypoglycemia clamps (5 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist)

N 13 ± 6 174 ± 38 40 ± 12 323 ± 50

D 52 ± 8 * 220 ± 19 93 ± 33 324 ± 29

D+SSTR2a 37 ± 6 226 ± 45 147 ± 40 282 ± 64

control experiments (no exogenous insulin administered)

Ctrl:N 50 ± 6 165 ± 34 31 ± 15 318 ± 11

Ctrl:D 45 ± 3 189 ± 54 110 ± 46 310 ± 82

Ctrl:D+SSTR2a 50 ± 4 245 ± 42 217 ± 65 290 ± 38

131

Table 4-3. Plasma insulin levels (ng/mL) during hypoglycemia clamps (10 U/kg or 5 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist). Data are represented as means ± SEM. *P<0.05 D vs N.

Time (min)

-60 0 60 180

Hypoglycemia clamps using 10 U/kg insulin and 3000 nmol/kg/h SSTR2 antagonist

N 1.0 ± 0.07 0.9 ± 0.07 141.5 ± 8.0 59.6 ± 7.4

D 0.5 ± 0.04 * 0.5 ± 0.07 * 93.2 ± 7.2 * 24.1 ± 2.9 *

D+SSTR2a 0.5 ± 0.1 0.7 ± 0.2 107.9 ± 14.1 26.3 ± 4.4

Hypoglycemia clamps using 5 U/kg insulin and 3000 nmol/kg/h SSTR2 antagonist

N 0.6 ± 0.4 0.6 ± 0.2 62.4 ± 13.2 5.2 ± 0.9

D 0.3 ± 0.2 0.2 ± 0.5 59.4 ± 17.8 1.5 ± 0.5

D+SSTR2a 0.3 ± 0.07 0.5 ± 0.2 58.6 ± 18.0 3.0 ± 1.5

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Table 4-4. Plasma counterregulatory hormone and insulin levels during control experiments in the absence of insulin (0 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist). “N/A” denotes time points at which samples for the particular hormone were not obtained. Data are represented as means ± SEM. *P<0.05 D vs N. †P<0.03 D vs D+SSTR2a.

Time (min)

-60 0 20 40 60 90 120 180

Plasma glucagon (pg/mL)

Ctrl:N 50 ± 6 70 ± 15 79 ± 24 64 ± 14 42 ± 12 47 ± 9 33 ± 9 39 ± 8

Ctrl:D 45 ± 3 50 ± 12 41 ± 13 43 ± 8 50 ± 10 51 ± 18 55 ± 10 59 ± 17

Ctrl:D+SSTR2a 50 ± 4 88 ± 18 101 ± 24 100 ± 22 † 97 ± 15 † 95 ± 11 † 80 ± 13 73 ± 13

Plasma corticosterone (ng/mL)

Ctrl:N 165 ± 31 210 ± 47 313 ± 36 294 ± 60 233 ± 56 195 ± 38 N/A 208 ± 29

Ctrl:D 205 ± 49 248 ± 75 351 ± 93 349 ± 84 335 ± 86 254 ± 74 180 ± 43

Ctrl:D+SSTR2a 273 ± 46 261 ± 53 316 ± 49 322 ± 55 331 ± 58 293 ± 50 225 ± 40

Plasma epinephrine (pg/mL)

Ctrl:N 31 ± 15 94 ± 19 89 ± 16 112 ± 19 85 ± 14 82 ± 25 N/A 45 ± 12

Ctrl:D 110 ± 46 85 ± 31 157 ± 43 116 ± 29 264 ± 112 382 ± 272 123 ± 49

Ctrl:D+SSTR2a 217 ± 65 180 ± 85 243 ± 156 385 ± 191 345 ± 149 371 ± 163 235 ± 111

Plasma norepinephrine (pg/mL)

Ctrl:N 356 ± 39 393 ± 45 428 ± 46 435 ± 88 353 ± 79 392 ± 78 N/A 293 ± 41

Ctrl:D 310 ± 82 292 ± 59 272 ± 31 310 ± 48 376 ± 96 301 ± 80 246 ± 108

Ctrl:D+SSTR2a 290 ± 38 238 ± 58 242 ± 100 381 ± 125 351 ± 76 315 ± 96 330 ± 105

Plasma insulin (ng/mL)

Ctrl:N 1.1 ± 0.2* 1.1 ± 0.1* N/A N/A 0.8 ± 0.1 N/A N/A 0.8 ± 0.1

Ctrl:D 0.7 ± 0.1 0.7 ± 0.1 0.7 ± 0.1 0.7 ± 0.1

Ctrl:D+SSTR2a 0.5 ± 0.1 0.7 ± 0.2 0.6 ± 0.2 0.6 ± 0.1

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Table 4-5. Quantitative mRNA expression of liver PEPCK1 and G6Pase following hypoglycemia clamp (10 U/kg insulin, 3000 nmol/kg/h SSTR2 antagonist) and control experiments in the absence of insulin (0 U/kg, 3000 nmol/kg/h). As per the 2-∆∆Ct method (526), gene expression was normalized to the control group, Ctrl:N. Data are represented as means ± SEM. *P<0.0001 Ctrl:N vs Ctrl:D. †P<0.05 Ctrl:D vs Ctrl:D+SSTR2a. ‡P<0.0001 Ctrl:N vs N. §P<0.01 Ctrl:D+SSTR2a vs D+SSTR2a.

mRNA expression

(fold-change relative to Ctrl:N group)

PEPCK1 G6Pase

Controls

Ctrl:N 1.000 ± 0.233 1.000 ± 0.301

Ctrl:D 0.395 ± 0.111 0.160 ± 0.041 *

Ctrl:D+SSTR2a 1.177 ± 0.280 0.499 ± 0.169 †

Hypoglycemia

N 1.039 ± 0.256 0.171 ± 0.049 ‡

D 1.293 ± 0.444 0.031 ± 0.008

D+SSTR2a 0.509 ± 0.183 0.033 ± 0.011 §

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Table 4-6. Metabolic parameters of non-diabetic rats treated with SSTR2 antagonist (N+SSTR2a) that underwent hypoglycemia clamp experiments using the different insulin (U/kg) and SSTR2 antagonist (nmol/kg/h) dosing combinations. Fed blood glucose, body weight, and food intake were measured daily, and the table below shows the mean of values averaged over the last week before the experimental day. Daily blood glucose measurements were obtained using a glucometer. Data are represented as means ± SEM.

fed blood glucose (mM)

body weight (g)

food intake (g)

N+SSTR2a

10 U/kg, 1500 nmol/kg/h (n=3)

5.7 ± 0.2 419 ± 9 19 ± 1

10 U/kg, 3000 nmol/kg/h (n=4)

5.2 ± 0.1 405 ± 22 19 ± 1

5 U/kg, 3000 nmol/kg/h (n=10)

5.3 ± 0.2 396 ± 21 16 ± 1

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Table 4-7. Pancreatic somatostatin protein levels (pmol/mg protein) and plasma somatostatin levels (pmol/mL) in all hypoglycemic rats. To assess the effects of diabetes and administration of SSTR2 antagonist on somatostatin, available data was combined for each treatment group, irrespective of doses of insulin or SSTR2 antagonist administered. Data are represented as means ± SEM. *P<0.04 D vs N. †P<0.005 D+SSTR2a vs N+SSTR2a. ‡P<0.03 D+SSTR2a vs N+SSTR2a. aP=0.08 D vs N. Additionally, statistical analysis using Two-Way ANOVA shows that there is an effect of diabetes, but not SSTR2 antagonist (P=0.51), on both pancreatic somatostatin (P<0.0004) and plasma somatostatin (P<0.004) values.

Pancreatic somatostatin

(pmol/mg protein)

Plasma somatostatin

(pg/mL)

N 85.3 ± 5.1 11.3 ± 2.4

D 121.1 ± 12.9 * 18.4 ± 2.3 a

N+SSTR2a 78.9 ± 7.8 12.0 ± 2.7

D+SSTR2a 130.4 ± 10.6 † 21.2 ± 2.2 ‡

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136

Chapter 5 Study 2: Amelioration of hypoglycemia

using SSTR2 antagonism

5 Study 2

5.1 Aims

Based on our prior observation that SSTR2 antagonism restored glucagon and

corticosterone counterregulation in acutely hypoglycemic diabetic rats, we wished to test

whether SSTR2 antagonism could ameliorate subsequent hypoglycemia by restoring blood

glucose levels to euglycemia, and whether this glycemic rescue could be attributed to

improved hormone counterregulation, despite exposure to recurrent hypoglycemia. Thus

the aim of the present study was to determine whether counterregulatory hormone

responses can be restored by SSTR2 antagonism in diabetic rats exposed to recurrent

hypoglycemia and if such improvements can lessen the severity of subsequent

hypoglycemia. Recurrent hypoglycemia is associated with marked impairments of

counterregulatory hormone responses. Glucagon counterregulation in diabetes is often

absent (165;166), thus it has not been shown to be further worsened by recurrent

hypoglycemia (203;227;264). However, there is evidence demonstrating markedly

diminished epinephrine counterregulation following recurrent hypoglycemia in diabetic

subjects (203;205;220;227;261;263;264). Cortisol responses following recurrent

hypoglycemia are less consistent, having been found unaltered (203;264) or decreased

(205;227) in diabetes. In the present study, diabetic rats were subjected to 5 prior

episodes of hypoglycemia (i.e. recurrent hypoglycemia treatment) in an attempt to create a

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model of markedly defective counterregulation. Our present results demonstrate that

SSTR2 antagonism promotes the recovery of hypoglycemia in recurrently hypoglycemic

diabetic rats and that improved glucagon counterregulation may be responsible, at least in

part, for this amelioration.

5.2 Experimental methods

5.2.1 Study design

Figure 5-1 illustrates the study design for the main protocol. As will be described in detail

below, two groups of STZ-diabetic rats were used in this study. Initial pilot studies

examined the effect of only 2 consecutive days of hypoglycemia (i.e. without prior recurrent

hypoglycemia treatment) and is presented in Figure 5-2 and Figure 5-3.

5.2.2 Recurrent hypoglycemia treatment

Rats were subjected to 5 episodes of recurrent hypoglycemia over 3 consecutive days (i.e. 2

episodes per day on the first two days and 1 episode on the third day). Rats were partially

fasted overnight prior to each day of recurrent hypoglycemia treatment. Partial fasting

consisted of 10-15 g of rat chow (i.e. approximately 25-40% of ad libitum consumption)

and free access to 5% sucrose water. Basal blood glucose was measured at the 0 min time

point, and regular insulin (20 U/kg; Humulin R, Eli Lilly, Indianapolis, IN) was injected

subcutaneously to induce each episode of hypoglycemia. Glucose infusions (50% dextrose)

were given at a variable rate to clamp glycemia at a target hypoglycemia of 3.0±0.5 mM.

Blood glucose was measured (Analox glucose analyzer, GMD-9D, Analox Instruments USA

Inc., Lunenburg, MA) in duplicate every 15 min for 180 min. During a rest period between

180 and 240 min, rats were given access to 5% sucrose water and intravenous glucose

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infusion to recover from hypoglycemia. One blood glucose sample was obtained during the

break to monitor glycemia. At 240 min, rats were given a second insulin injection as above

and remained at hypoglycemic until 420 min. Food and sucrose water were fed to aid

recovery after hypoglycemia treatment.

5.2.3 Partial overnight fasting prior to hypoglycemia experiments

Rats were partially fasted overnight prior to each experimental day. This food restriction

consisted of 10-15 g of rat chow (i.e. approximately 25-40% of ad libitum consumption)

given overnight when rats are awake. Fasting rats in this manner was necessary for three

purposes: (i) it allowed target hypoglycemia to be achieved more easily during recurrent

hypoglycemia treatment as compared to non-fasted conditions; (ii) it allowed rats to

preserve some glycogen stores to facilitate recovery from hypoglycemia during

Experimental Days 1 and 2; (iii) and it allowed for better control of body weight and initial

glycemia, which was important since insulin dosing was dependent on these variables and

we wished to keep the amount of insulin administered consistent between the 2

experimental days. Limitations to this fasting method included an inability to precisely

determine the time at which rats ate their rationed chow; we narrowed the window of

rationed chow availability to between 22:00 (when chow was weighed and rationed) and

09:00 (when rat cages where changed, any uneaten food removed, and 5% sucrose water

replaced with regular water).

5.2.4 Hypoglycemia on Experimental Days 1 and 2

Following recurrent hypoglycemia treatment, each rat subsequently underwent 2 days of

experimentation. The protocol for experimental day 1 (ExptD1) and experimental day 2

(Expt2) is summarized in Figure 5-1. On the morning of the experiments, rats were

weighed and had their cages changed, connected to infusion lines, and acclimatized for a

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minimum of 2 h prior to the start of experimentation. Basal blood samples for glucose and

hormones were taken at the start of the experiment (i.e. t=-60 min) from freely moving,

conscious rats. Extreme care was taken to avoid stresses to the rats during the

experiments since it is well known that even minimal disturbances caused by sounds,

scents, and vibrations can increase the release of the stress hormones measured. On

ExptD1, 0.9% saline infusion (1 mL/h) was started after basal samples were obtained at the

-60 min time point. Blood glucose was measured using a glucose analyzer in duplicate at

times -60, -40, -20, 0, and every 10 min thereafter until 240 min. Blood samples were

obtained from the carotid catheter at regular time intervals throughout the experiment for

measurement of plasma hormone levels. After plasma was removed, packed red blood cells

were re-suspended in heparinized saline (10 USP U/mL) containing 1% bovine serum

albumin and re-infused into the rat. After blood samples were obtained at t=0, an

intravenous insulin bolus (10 U/kg) was administered, and a primed intravenous insulin

infusion (50 mU/kg/min) was commenced and terminated specifically for each rat at the

experimenter’s discretion when target hypoglycemia (<4.0 mM) was approached. Infusions

were delivered via digital pumps (Harvard Apparatus PHD 22/2000 syringe pumps,

Holliston, MA), and both the volume of insulin infused and the time at which the infusion

was stopped were recorded. The purpose of Experimental Day 1 was to precisely determine

the minimal amount of insulin necessary to induce hypoglycemia in a particular rat.

Determining the insulin dosage specifically for each rat was necessary since the insulin

sensitivities of these diabetic rats varied. No glucose infusions or SSTR2 antagonist were

given on the experimental days.

On ExptD2, rats were divided into the SSTR2 antagonist treatment group (saline-SSTR2a,

n=13) or control group (saline-saline, n=7). In each rat, the same insulin administration

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regime was used as ExptD1. It is important to emphasize that: (i) the same rat was used in

both ExptD1 and ExptD2 for comparisons; (ii) the SSTR2 antagonist treatment group and

the control group represent two different groups of rats. In the treatment group, SSTR2

antagonist infusion (3000 nmol/kg/min at 1 mL/h) was commenced at t=-60 min and

continued for 5-h duration of the experiment to determine the effect of SSTR2 antagonism

on the extent (i.e. depth and duration) of hypoglycemia. This dose of SSTR2 antagonist

was chosen because it normalized glucose and corticosterone responses to acute

hypoglycemia in diabetic rats and did not elicit hyperglycemia or marked elevations in

counterregulatory during an absence of hypoglycemia (Study 1). In the control group,

saline was infused without SSTR2 antagonist to control for the repeated bout of

hypoglycemia (i.e. saline infusions on both ExptD1 and ExptD2). Body weight, initial

glycemia, and overnight food intake were controlled for both days in all rats. At the end of

240 min on ExptD2, rats were quickly euthanized by decapitation. Rats that required

glucose infusion due to severe hypoglycemia (i.e. < 1.6 mM) were excluded from the data

presented.

5.3 Results

5.3.1 Pilot Studies: How the present method was established

Initial pilot studies examined the effect of only 2 consecutive days of hypoglycemia (i.e.

without prior recurrent hypoglycemia treatment) in STZ-diabetic rats in which insulin was

administered subcutaneously with repeated staggered injections (Figure 5-2; n=4; average

of 4.8 U insulin injected) or intravenously via bolus and infusion (Figure 5-3; n=3; average

of 4.9 U insulin administered). Repeated, staggered subcutaneous injections proved to be

problematic since: i) unwanted stress was caused, and ii) precise dosing was difficult to

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administer. In both studies, as in the data presented below, a minimal amount of insulin to

induce hypoglycemia was administered on the first day, the same amount of insulin was

given on the second day to the same rat, and no glucose infusion was given so that

recovery could be assessed. In these pilot studies, rats were overnight fasted. In both

studies, one prior bout of hypoglycemia the day before greatly worsened hypoglycemia

induced with the same amount of insulin on the next consecutive day as was apparent both

by the measurement of blood glucose levels (Figure 5-2A, Figure 5-3A) as well as the

observable behavioural condition of the rats (i.e. immobilization, and in many rats, severe

twitches, which may have been seizures). One antecedent bout of hypoglycemia was

sufficient to markedly diminish glucagon responses to subsequent hypoglycemia (Figure

5-2B, Figure 5-3B). We reasoned that a protocol that involved recurrent hypoglycemia

treatment in a clamped setting prior to experimentation would be useful for the following

reasons: i) from a practical standpoint, several prior repeated episodes of hypoglycemia

would help to minimize the glycemic variation between subsequent bouts of hypoglycemia

(i.e. we hypothesized a habituation effect); ii) it would allow us to evaluate the effect of

SSTR2 antagonist in a established model of severe counterregulatory impairment; and iii) as

we hypothesized that epinephrine counterregulation would be diminished in recurrently

hypoglycemic diabetic rats (unlike in our acute hypoglycemia setting in Study 1), it would

allow us to evaluate the effect of SSTR2 antagonist on epinephrine counterregulation. Due

to the marked effect of one antecedent bout of hypoglycemia on impairing hypoglycemic

recovery, we also decided to partially fast, instead of fully fast, rats overnight.

5.3.2 Daily blood glucose, body weight, and food intake

Metabolic parameters such as fed morning blood glucose, body weight, and food intake

were measured daily to monitor the health of the rats and to ensure that diabetic rats were

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of a similar diabetic state leading up to recurrent hypoglycemia treatment. In the week

prior to recurrent hypoglycemia, fed morning hyperglycemia, body weight, and daily food

intake were the same in rats that would later be divided into SSTR2 antagonist treatment

and controls groups for hypoglycemia experiments (Table 5-1).

5.3.3 Glycemia during recurrent hypoglycemia treatment

Both groups achieved 5 similar episodes of recurrent hypoglycemia over 3 days (3.0±0.5

mM for an average of 90 min per episode; Figure 5-4).

5.3.4 Basal blood glucose and plasma hormone levels after recurrent hypoglycemia treatment

On the morning of Experimental Days 1 and 2, body weight and initial glycemia did not

differ between rats (Table 5-2). Thus, all rats had similar metabolic starting points following

recurrent hypoglycemia, which allowed for baseline control conditions. Basal plasma

glucagon, epinephrine, norepinephrine, and insulin were also similar in all rats (Table 5-3).

Oddly, basal corticosterone of ExptD2-SSTR2a was lower compared with its respective

ExptD1 (P<0.03), whereas basal corticosterone was unchanged in controls (Table 5-3).

This is unusual because this change cannot be attributed to any changes in the metabolic

parameters measured, to the levels of basal hormones measured, to recurrent

hypoglycemia, or to SSTR2 antagonist since this was before the antagonist was

administered.

5.3.5 Blood glucose and plasma insulin levels during Experimental Days 1 and 2

On ExptD1, insulin induced a similar rate of decline in blood glucose levels to <4.0 mM by

90 min in both treatment (Figure 5-5) and control (Figure 5-6) groups. The hypoglycemic

nadirs were likewise the same in both treatment (Figure 5-7A) and control groups (Figure

5-7B) on ExptD1. The control group had a longer duration of hypoglycemia compared to

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the treatment group. When SSTR2 antagonist was administered on ExptD2 with the same

insulin administration regime (i.e. same bolus dose and same infusion dose, rate, and

timing of administration) as ExptD1, a similar glycemic decline from hyperglycemia ensued,

but the depth and duration of hypoglycemia was significantly less when SSTR2 antagonist

was given (Figure 5-7A). This demonstrates that SSTR2 antagonism does not affect insulin

sensitivity or its glycemia lowering effect. SSTR2a rats recovered from hypoglycemia more

quickly, and the hypoglycemic nadir was less severe in ExptD2-SSTR2a (3.7±0.3 mM) than

on the preceding day (2.9±0.1 mM; P<0.01). The extent of hypoglycemia, calculated as

the area under the curve below 4.0 mM, was significantly reduced in the SSTR2 antagonist

treatment group (P<0.001; Figure 5-7A). Analysis of time points after glycemia in both

groups fell below 7.0 mM (i.e. t=50-240min) or 4.0 mM (i.e. t=110-240 min) with Repeated

Measures ANOVA revealed a significant effect of SSTR2 antagonist (P<0.02 and P<0.005,

respectively). Without the SSTR2 antagonist, there was no improvement in hypoglycemic

recovery (Figure 5-7B). Controls had similar hypoglycemia on both days (nadirs: 2.8±0.4

mM).

As we carefully endeavoured, similar amounts of insulin (via bolus, infusion, and total

amount given) were administered to both groups on both days (Table 5-2). Giving the

same amount of insulin on both experimental days within a group was important so that any

changes observed with glycemia would not be attributed to the amount of insulin

administered. This resulted in similar plasma insulin profiles for both groups (Table 5-4).

5.3.6 Counterregulatory hormones levels during Experimental Days 1 and 2

Following recurrent hypoglycemia, glucagon responses on Experimental Day 1 in both

groups (Figure 5-8) were modest. Figure 5-8A shows all rats that received SSTR2

antagonist on ExptD2 and their respective glucagon responses on ExptD1. Figure 5-8B

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omits six rats that did not experience hypoglycemia as a result of their SSTR2 antagonist

treatment on ExptD2 and their respective glucagon responses on ExptD1 (i.e. Figure 5-8B

shows data for both experimental days of rats whose blood glucose levels dipped below the

normal physiological threshold of 4.0 mM in response to insulin administration despite

SSTR2 antagonist treatment on ExptD2). Figure 5-8C shows the glucagon responses of

control rats (i.e. no SSTR2 antagonist administered on ExptD2). It should be noted that

none of the ExptD2-control rats had hypoglycemia prevented. SSTR2 antagonist

significantly increased plasma glucagon on ExptD2 by 3-fold as compared to ExptD1

(P<0.04, Figure 5-8A) and had a strong tendency to improve plasma glucagon by 2.2-fold in

rats that still experienced hypoglycemia (P=0.05, Figure 5-8B), whereas the glucagon

response in controls remained markedly attenuated and exhibited a trend to be decreased,

although this was not statistically significant (Figure 5-8C).

Epinephrine responses were significantly lower in ExptD2-SSTR2a rats than on their

respective ExptD1 (Figure 5-9A). We attribute this attenuated epinephrine response to the

improvement in hypoglycemic recovery on ExptD2 due to SSTR2 antagonism; since many of

these rats did not experience marked hypoglycemia, there was no need for them to respond

with robust epinephrine counterregulation. When epinephrine responses were analyzed

having omitted the six rats in which hypoglycemia was prevented on ExptD2 due to

administration of SSTR2 antagonist, it is observed that SSTR2 antagonist preserves

epinephrine counterregulation to recurrent hypoglycemia (Figure 5-9B). In contrast,

ExptD2-controls had a significantly blunted epinephrine counterregulation (38% reduction)

compared to their respective responses on ExptD1 (P<0.02; Figure 5-9C). As mentioned

above, none of the ExptD2-control rats had hypoglycemia prevented. This suggests that

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SSTR2 antagonism can help preserve the epinephrine response in recurrently hypoglycemic

rats when exposed to subsequent hypoglycemia.

Corticosterone responses likewise were decreased in ExptD2-controls by 46% compared to

their responses on ExptD1 (P<0.04, Figure 5-10C). However, treatment in ExptD2-SSTR2a

preserved the corticosterone response to hypoglycemia following recurrent hypoglycemia

(Figure 5-10A). Omission of the six rats in which hypoglycemia was prevented did not

significantly alter the corticosterone profile of the ExptD2-SSTR2a group, although these

rats demonstrated a strong tendency for increased corticosterone counterregulation

(P=0.05; Figure 5-10B).

Despite similar metabolic profiles (i.e. body weight, severity of diabetic hyperglycemia at

baseline), similar recurrent hypoglycemia treatment, and similar insulin administration and

plasma levels, ExptD1 epinephrine and corticosterone responses to hypoglycemia were

greater in controls than in the treatment group. Similar counterregulatory hormone

responses following recurrent hypoglycemia were expected on ExptD1, which occurred

before SSTR2 antagonist treatment, since rats had similar metabolic profiles and were

treated in the same way up until this point. We attribute the larger epinephrine and

corticosterone responses of the ExptD1 control group to a greater extent of hypoglycemia

(Figure 5-7B) as compared with ExptD1 in the treatment group (Figure 5-7A). This

supports the idea that different rats have different insulin sensitivities; given the same

amount of insulin, some rats may be more susceptible to hypoglycemia than others. Thus,

it was important that we used the method of comparing the same rat to the effects of

recurrent hypoglycemia on ExptD1 and ExptD2, with (i.e. treatment group) and without (i.e.

control group) SSTR2 antagonist.

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Norepinephrine levels were similar on both experimental days in each of the groups (Figure

5-11).

Figure 5-12 compares the peak incremental hormone responses to acute hypoglycemia from

Study 1 and recurrent hypoglycemia from Study 2 in diabetic control rats and diabetic rats

treated with SSTR2 antagonist and will be discussed below.

5.4 Discussion

5.4.1 Amelioration of hypoglycemia with SSTR2 antagonism

We demonstrate that SSTR2 antagonism promotes the recovery of hypoglycemia by

reducing both the depth and duration of hypoglycemia in recurrently hypoglycemic diabetic

rats. Whereas control animals subjected to recurrent hypoglycemia had glycemic nadirs of

2.8±0.4 mM on both experimental days, rats given SSTR2 antagonist had a significantly

improved glycemic profile as demonstrated by a shorter duration of hypoglycemia and an

elevated glycemic nadir of 3.7±0.3 mM. Lessening the hypoglycemic nadir is important to

preserve brain function. An early study demonstrated that hypoglycemia at 3.3 mM

reduced cognitive function as compared with euglycemia in diabetic subjects (620). Blood-

to-brain glucose transport is a direct function of the arterial plasma glucose concentration,

and during hypoglycemia blood-to-brain glucose transport becomes limiting to brain glucose

metabolism and, therefore, function (4). The American Diabetes Association Workgroup on

Hypoglycemia has emphasized not only the inherent danger of the nadir glucose

concentration and the duration of hypoglycemia but also of the frequency of hypoglycemic

events (1). Some (151;621;622) have challenged the substantiality of defining

hypoglycemia as 3.9 mM, as published by the American Diabetes Association Workgroup

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on Hypoglycemia (1). Amiel et al. suggest that glycemic excursions in the range of 3.5–4.0

mM, provided that further glycemic decline is prevented, is of little clinical significance

(151). Swinnen et al. report that defining hypoglycemia at a higher glucose level of 3.9

mM overestimates the prevalence of asymptomatic episodes of hypoglycemia and suggest a

lower glycemic cut-off (622). Frier suggests that a cut-off level of 3.5 mM or lower to

define the onset of hypoglycemia would be clinically appropriate since cognitive function is

not impaired, and symptoms are often not generated, until glycemia approaches 3.2 mM

(621). If this were indeed the case, then our present data would illustrate that SSTR2

antagonist treatment could not only facilitate recovery from low blood glucose levels but

also prevent the onset of hypoglycemia. However, it should be noted that a more

conservative definition of hypoglycemia (i.e. at the biochemical level of 3.9 mM) has its

merits as rebutted by Cryer (623). Hypoglycemia, even when unaccompanied by

symptoms, leads to defective glucose counterregulation and eventually hypoglycemia

unawareness (63;168). Since these episodes substantially increase the risk of subsequent

hypoglycemia, all hypoglycemic events can harm the individual with diabetes in the short or

long term (1). It has been suggested that delayed recovery of hypoglycemia may

frequently occur in type 1 diabetic individuals in which deficient epinephrine and glucagon

counterregulation results in impaired hepatic glucose release (225). Thus, developing a

means to promote hypoglycemic recovery is of vital importance.

Hypoglycemia unawareness is the development of neuroglycopenia without appropriate prior

autonomic warning symptoms (232;624) and contributes to the increased risk of recurrent

hypoglycemia (63). In type 1 diabetic patients, avoidance of hypoglycemia could restore

hypoglycemia awareness. Improved symptom scores, increased glycemic thresholds for

responses, and/or improved epinephrine counterregulation were observed after 2 weeks

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(625), 3 months (625;626), or 4 months (238;241). However, the restoration of awareness

would often come at the cost of less strict glycemic control as measured by increased

hemoglobin A1C levels (241;625;626). Thus, our findings suggest a means by which

hypoglycemia can be ameliorated without compromising glycemic management.

So how might SSTR2 antagonism facilitate the recovery of hypoglycemia? We demonstrate

that facilitated recovery from hypoglycemia using SSTR2 antagonist is associated with an

improvements in glucagon, epinephrine, and corticosterone responses to recurrent

hypoglycemia. As will be discussed in section 5.4.2, improvements of hormone

counterregulation could increase glucose production during hypoglycemia to help restore

euglycemia, thus ameliorating hypoglycemia via SSTR2 antagonism.

It has been demonstrated that central administration of somatostatin has no effect on

plasma glucose levels under basal conditions. However, when somatostatin was

administered into the central nervous system in conjunction with -endorphin or bombesin,

the hyperglycemia that would typically ensue from each of these substances was abolished

(627;628). An elevation of blood glucose levels is achieved by increasing glucose

production and decreasing glucose clearance. From Study 1, 4-h infusion of SSTR2

antagonist in diabetic rats during hypoglycemia yielded no changes in growth hormone

levels as compared with saline infused controls, thus it is unlikely that growth hormone-

induced hepatic glucose output contributed to improved hypoglycemic recovery. The

inability of somatostatin to directly affect glucose turnover both in vivo in dogs (629) and in

vitro in rat hepatocytes (616) has been reported by much earlier studies by Cherrington et

al. In dogs, infusion of somatostatin with intraportal replacement co-infusions insulin and

glucagon did not affect glucose production, which suggests that somatostatin may affect

glucose production indirectly via its effects on insulin and glucagon, but not directly (629).

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However, the lack of effect of somatostatin reported in these studies was during the basal,

non-stimulated state and is does not necessarily represent conditions of hepatic stimulation.

In perfused rat liver, somatostatin inhibited glucagon-stimulated, but not basal, hepatic

glucose output (630). Since somatostatin also blunted cAMP-induced glucose output, the

authors suggest the effect of somatostatin to suppress stimulated hepatic glucose release is

mediated via cAMP (630), which is consistent with a known mechanism of action of

somatostatin. Furthermore, in isolated hepatocytes, somatostatin inhibited both glucagon-

stimulated glycogenolysis and gluconeogenesis by 40-50%, but somatostatin had no effect

on epinephrine-stimulated glycogenolysis (631). At present, however, there is no known

direct effect of SSTR2 antagonism on hepatic glucose production. SSTR1 (271) and SSTR3

(373), but not SSTR2 (418;632), have been detected on hepatocytes, and the SSTR2

antagonist has no reactivity with SSTR1 and 10-fold less binding affinity with SSTR3 (509).

We demonstrate that SSTR2 antagonism did not affect the mRNA expression of

gluconeogenic enzyme PEPCK in diabetic rats during basal conditions or in response to

hypoglycemia (Study 1). Similarly, SSTR2 antagonist did not affect expression of G6Pase

following hypoglycemia but increased G6Pase expression during basal conditions (Study 1).

In another study, however, SSTR2-deficient mice were shown to have increased mRNA

expression of PEPCK, but not G6Pase (592).

It has been postulated that an increase in glucose clearance may be attributed to some

extrapancreatic effect of somatostatin (633). In one study, somatostatin infusion did not

increase whole-body glucose disposal in non-diabetic humans during a hyperinsulinemic-

euglycemic clamp (298). In contrast, a direct effect of somatostatin to enhance insulin-

stimulated muscle glucose uptake, but not basal muscle glucose uptake, was demonstrated

in non-diabetic humans (634). During a hyperinsulinemic-euglycemic clamp, local forearm

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perfusion (via the brachial artery) with somatostatin elicited a significant 55% increase in

insulin-stimulated forearm glucose uptake as compared with systemic venous somatostatin

infusion without affecting the exogenous glucose requirement or plasma insulin, glucagon,

or growth hormone levels. The authors concluded that somatostatin may increase insulin-

stimulated glucose uptake in muscle via local mechanisms (634). Thus, it is plausible that

inhibition of somatostatin in the present study could contribute to reduced muscle glucose

uptake during hypoglycemia, thus increasing blood glucose levels. Evidence of SSTR

subtypes on skeletal muscle is scarce, but one group has reported detection of SSTR2,

SSTR3, and SSTR4 mRNA in rat skeletal muscle (635) while another group has evidence of

SSTR5 mRNA in skeletal muscle (636).

5.4.2 SSTR2 antagonism improves counterregulation following recurrent hypoglycemia

Recurrent hypoglycemia is well known to impair counterregulatory hormone responses in

both non-diabetic (227;265;637-640) and diabetic (203;205;220;227;261;263;264)

subjects. We also demonstrate that the peak incremental glucagon (P<0.005) and

epinephrine (P<0.001) responses to subsequent hypoglycemia are blunted in recurrently

hypoglycemic rats as compared to rats subject to acute hypoglycemia (Figure 5-12).

5.4.2.1 SSTR2 antagonism increases glucagon counterregulation.

Presently, we provide evidence that SSTR2 antagonism increases plasma glucagon levels in

recurrently hypoglycemic diabetic rats. In contrast, the glucagon response to recurrent

hypoglycemia was absent in control rats without SSTR2 antagonist. In the present study,

we were unable to collect and analyze pancreata from our experimental animals. However,

we previously observed increased pancreatic prosomatostatin mRNA expression in diabetic

rats as compared to non-diabetic rats (205). After 7 episodes of recurrent hypoglycemia in

diabetic rats, the expression of somatostatin was slightly but not significantly decreased

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compared to diabetic controls without recurrent hypoglycemia but remained partially

elevated in as compared with non-diabetic controls, which indicates partial normalization of

pancreatic somatostatin expression following recurrent hypoglycemia (205). In contrast, 7

recurrent episodes of recurrent hyperinsulinemia, per se, decreased but did not fully

normalize pancreatic somatostatin mRNA, which demonstrates that repeated insulin

administration, and not repeated hypoglycemia, may help to partially normalize pancreatic

somatostatin expression. In these diabetic rats that received recurrent hyperinsulinemic

treatment, with or without recurrent hypoglycemia, and had partially normalized pancreatic

somatostatin expression, glucagon responses to subsequent hypoglycemia was improved as

compared with acutely hypoglycemic diabetic controls but still impaired compared to non-

diabetic controls (205). Thus, as with acute hypoglycemia, increased pancreatic

somatostatin may play a role in impairing glucagon release in response to repeated

hypoglycemia.

It should be noted that although SSTR2 antagonism improved glucagon responsiveness in

recurrently hypoglycemic rats by 3-fold, this peak incremental response was much less than

the glucagon response in SSTR2 antagonist-treated diabetic rats during acute hypoglycemia

from Study 1 (30±10 vs. 228±57 pg/mL; Figure 5-12A). In a separate study we previously

demonstrated that chronic, modest insulin treatment in STZ-diabetic rats does not

normalize glucagon counterregulation following recurrent hypoglycemia (641). Thus,

recurrent hypoglycemia still overwhelmingly attenuates glucagon counterregulation,

irrespective of insulin treatment, and we suggest that SSTR2 antagonism may offer

significant improvement.

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5.4.2.2 SSTR2 antagonism preserves epinephrine counterregulation.

A very important defect of counterregulation in many diabetic individuals is the poor

response of epinephrine, especially in conjunction with an absent glucagon response and

hypoglycemia unawareness (3). Previously, we observed a deficiency of the epinephrine

response to recurrent hypoglycemia in STZ-diabetic rats (205), and this defect was

attributed, in part, to deficiencies in the mRNA expression of two key catecholamine-

synthesizing enzymes (220). Furthermore, it has also been shown that although BB

diabetic rats had normal total catecholamine content in the adrenal medulla, they showed a

hypofunctional adrenomedullary chromaffin cell release of epinephrine (642). A central

effect of somatostatin on regulating epinephrine cannot be ruled out. Prior studies have

demonstrated that analogues of somatostatin inhibit the plasma catecholamine response to

central glucopenia and to central administration of carbachol and bombesin (528;643).

We presently hypothesized that antagonizing somatostatin may play a role in increasing

secretion of adrenomedullary epinephrine in response to recurrent hypoglycemia. In our

prior acute hypoglycemia study (Study 1), we observed no effect of the SSTR2 antagonist

on epinephrine levels in diabetic rats, which had the same epinephrine response as non-

diabetic controls. We postulated that no further enhancement by SSTR2 antagonist could

be observed in those rats because the maximal physiological epinephrine response was

already achieved. As compared to our acute hypoglycemia study, peak increments in

plasma epinephrine levels following recurrent hypoglycemia were markedly reduced (acute

diabetic control: 3845±358 vs. recurrently hypoglycemic diabetic control: 1013±270

pg/mL), demonstrating that our recurrent hypoglycemia treatment generated a model of

defective counterregulation. The findings from the present study suggest that: (i) recurrent

hypoglycemia further decreased epinephrine levels in controls; (ii) when hypoglycemia is

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prevented, administration of SSTR2 antagonist does not increase plasma epinephrine levels;

but that (iii) SSTR2 antagonism helps to preserve the epinephrine response from further

decrement in rats whose glycemia dips below the normal physiological range, which may aid

in the promoted recovery from hypoglycemia.

SSTR2 has been widely identified in the adrenal medulla (390;397-399). It has been shown

that somatostatin inhibits acetylcholine-stimulated release of epinephrine from the adrenal

medulla (644;645). This is the mechanism whereby epinephrine is released during

hypoglycemia and other stressors (646). Somatostatin is postulated to inhibit receptor-

coupled signalling initiated by acetylcholine-nicotinic receptor binding, subsequent

membrane depolarization, and intracellular calcium increase (647-649), resulting in the

consequent inhibition of adrenomedullary epinephrine secretion from the adrenal medulla.

Furthermore it has been shown that although diabetic rats had normal catecholamine

content in the adrenal medulla, a defect in the secretory mechanism of epinephrine from

adrenomedullary chromaffin cells was observed (642). Therefore, we hypothesize that

administration of the SSTR2 antagonist may improve or normalize the epinephrine response

to hypoglycemia in diabetic rats. Co-storage and co-secretion of somatostatin with

catecholamines in chromaffin cells of the adrenal medulla granules have been demonstrated

(650). Thus, somatostatin may have the physiologic role of a negative feedback mechanism

for catecholamine release in the adrenal medulla (647). In addition, it is well known that

somatostatin administered directly in the brain reduces stimulated epinephrine responses.

Intracerebroventricular administration of somatostatin or its analogs reduces stress-induced

plasma epinephrine levels in non-diabetic rats and non-diabetic and diabetic dogs

(169;528;643;651;652). Thus, in addition to the direct effect of SSTR2 blockade at the

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adrenal medulla, it is also possible that inhibition of SSTR2 in the brain may indirectly

increase plasma epinephrine responses to hypoglycemia.

An improvement in the epinephrine response might also contribute to the amelioration of

hypoglycemia as observed in this study. It has been reported that in many type 2 diabetic

individuals with insulin resistance, the lipolytic effects of epinephrine outweigh the effects of

insulin on adipose tissue (186). Plasma free fatty acids increase in response to

hypoglycemia in type 2 diabetes (167;183;186) but not in type 1 diabetes (653).

Epinephrine secretion during hypoglycemia may therefore have a greater protective effect in

insulin-resistant subjects by promoting metabolic substrate release rather than storage.

Epinephrine also stimulates release of glucose from the kidney, and, in subjects who have a

deficient glucagon response to hypoglycemia, this has been suggested to compensate for

their impaired hepatic glucose output (213).

Norepinephrine counterregulation was not affected by recurrent hypoglycemia; peak

incremental norepinephrine responses were similar in diabetic control rats following acute

and recurrent hypoglycemia (565±43 vs. 604±129 pg/mL, respectively). SSTR2

antagonism had no effect on plasma norepinephrine levels in recurrently hypoglycemic rats.

This finding is consistent with the report that intracerebroventricular administration of

somatostatin or its analogs does not reduce stress-induced plasma elevations of plasma

norepinephrine (654).

5.4.2.3 SSTR2 antagonism preserves corticosterone counterregulation.

The glucocorticoid response to hypoglycemia plays a role in reinstating euglycemia, is

important during prolonged hypoglycemia, but may not play a critical role in rapid

counterregulation if other responses are intact (3). We have demonstrated that SSTR2

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antagonism can improve the corticosterone response to acute hypoglycemia in diabetic rats

(Study 1). In this study, we report that SSTR2 antagonist preserves the corticosterone

response to recurrent hypoglycemia whereas rats exposed to recurrent hypoglycemia

without SSTR2 antagonism display attenuated corticosterone counterregulation. Previously,

we demonstrated that insulin-treated STZ-diabetic rats had an absent corticosterone

response following recurrent hypoglycemia (641). We attributed this defect mainly to a lack

of suppression of hippocampal mineralocorticoid receptor mRNA expression and a

consequent lack of increase of hypothalamic CRH mRNA expression in response to

hypoglycemia (655). Intracerebroventricular administration of somatostatin blunts stress-

induced plasma cortisol levels in non-diabetic and diabetic dogs (169;528). Furthermore,

central administration of somatostatin inhibits pituitary release of ACTH (656), which

stimulates adrenocortical release of corticosterone. SSTR2 have been localized in several

brain regions and specifically in areas pertaining to HPA activity – the hypothalamic

paraventricular nucleus (420) and ACTH cells in the anterior pituitary (561). Interestingly,

somatostatin inhibits CRH and ACTH release (271;294;657;658), which regulate

corticosterone secretion. At present, the precise mechanism of somatostatin on its

regulation of the HPA axis in unknown, but we postulate that somatostatin may play a role

in regulating the stress response to hypoglycemia.

5.4.3 Discussion of other mechanisms for reduced counterregulation following recurrent hypoglycemia

Hypoglycemia diminishes hormone, symptom, and cognitive responses to subsequent

hypoglycemia in type 1 diabetic (203;223;226), type 2 diabetic (168), and non-diabetic

individuals. Recurrent hypoglycemia alters the thresholds for these responses such that

lower blood glucose levels must be reached before counterregulation is initiated (4).

Notably, the epinephrine response to hypoglycemia is diminished by recurrent

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hypoglycemia, which plays a role both in defective glucose counterregulation due to absent

glucagon counterregulation and hypoglycemia unawareness. Taken together, recurrent

hypoglycemia and the associated impairment of epinephrine counterregulation substantially

contribute to hypoglycemia-associated autonomic failure (HAAF) (63). Several different

mechanisms have been hypothesized to contribute to these impairments associated with

recurrent hypoglycemia.

Several studies have demonstrated that recurrent hypoglycemia alters glucose sensing in

the brain (193). The VMH contains glucose-sensing neurons (24;40;659) and also is a

critical region for the initiation of hypoglycemic counterregulation (34-36). Glucose-

inhibited (GI) neurons in the VMH increase their firing rate when extracellular glucose levels

decrease (40). Recurrent hypoglycemia impairs the response of VMH GI neurons to low

glucose conditions and lowers the glucose threshold at which neurons are activated

(263;660). Chan et al. have demonstrated that increased VMH GABAergic tone suppresses

counterregulation following recurrent hypoglycemia (661). This effect of increased GABA

inhibitory tone resulting from recurrent hypoglycemia is believed to be mediated via

glucose-excitatory (GE) neurons in the VMH. A recent hypothesis by McCrimmon posits that

recurrent hypoglycemia alters the balance between the 2 subpopulations of glucose sensing

neurons: the GI neurons and GE neurons. It is hypothesized that GI neurons, which act to

amplify counterregulation, are less likely to be active and/or GE neurons, which act to

suppress counterregulation, are more likely to be active following recurrent hypoglycemia

(193). This hypothesis is consistent with the studies mentioned above which demonstrate

that in the VMH, recurrent hypoglycemia increased GABAergic inhibitory tone (which is

postulated to be GE neuron-mediated) and decreased AMPK activity (which is postulated to

be GI neuron-mediated) (193). Thus, the decreased counterregulatory hormone responses

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observed in the recurrently hypoglycemic rats of the present study may be due to

recurrent-hypoglycemia induced alterations in brain glucose sensing.

In addition to the VMH, other brain areas have also been implicated in regulating

counterregulation to hypoglycemia. Integration of behavioural responses to hypoglycemia

is associated with brain networks that include the amygdala, brain stem, orbitofrontal cortex

(cognitive decision-making), and ventral striatum (reward centre) (662). Hypoglycemia

unaware subjects, who also had decreased epinephrine counterregulation, demonstrated

reduced responses in these networks, and it is hypothesized that habituation of these

behavioural responses may contribute to hypoglycemia unawareness (662). Furthermore,

the dorsal midline thalamus has been demonstrated to play an inhibitory role following

recurrent hypoglycemia and contribute to HAAF by reducing both symptom and epinephrine

responses to subsequent hypoglycemia (663). Thus, the counterregulatory defects

observed following recurrent hypoglycemia are regulated by complex neural integration of

both glucose sensing and afferent neural pathways that initiate counterregulatory

responses.

Another hypothesized mechanism relates to increased transport of glucose or other fuels to

the brain following recurrent hypoglycemia. When the brain fails to sense a local decrease

in glucose concentrations as a result of increased blood-to-brain transport of glucose or

alternate fuels, the brain reduces its initiation of counterregulatory hormone responses

(664;665). Boyle et al. have demonstrated adaptations that maintain brain glucose uptake

at normal levels following recurrent hypoglycemia and in type 1 diabetic individuals, thereby

reducing neuroendocrine counterregulatory responses (178;666). Prolonged, chronic

hypoglycemia of at least 3 days increases glucose transporter (GLUT)-1 mRNA and protein

at the blood-brain-barrier and glucose uptake in rats (177;667;668). However, this

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hypothesis has been challenged in light of evidence that demonstrated that 24 h of

hypoglycemia in humans blunted counterregulation to subsequent hypoglycemia but did not

affect glucose transport, glucose metabolism, or blood flow in the brain (669).

Furthermore, recurrent hypoglycemia of a shorter duration blunts hormone

counterregulation, but increases of local glucose concentrations in various brain regions

during subsequent hypoglycemia have not been observed. Thus, recurrent intermittent

hypoglycemia may not be a potent enough stimulus to cause alterations in brain glucose

transport that could be responsible for the defects of hormone counterregulation to

subsequent hypoglycemia presently observed.

5.4.4 Why might SSTR2 antagonism not have as great of an effect on recurrent hypoglycemia as acute hypoglycemia?

In Study 1, we consistently observed robust restoration of glucagon and corticosterone

counterregulation to acute hypoglycemia in diabetic rats with SSTR2 antagonism. Although

glucagon, corticosterone, and epinephrine responses in Study 2 are shown to be

significantly increased or preserved, the magnitudes of these improvements with SSTR2

antagonist treatment are not as profound following recurrent hypoglycemia. One possibility

to explain this difference of the glucagon response is that STZ-diabetic rats are more insulin

resistant than human type 1 diabetes, and thus much greater doses of insulin must be used

to induce hypoglycemia in these animal experiments than is usually used in insulin-

treatment of human diabetes. Thus following recurrent hypoglycemia, the sensitivity of

diabetic pancreatic -cells to the suppressive effects of exogenously administered insulin

becomes so overwhelming that disinhibition of somatostatin’s effect on the -cell is

ineffective at stimulating glucagon release to hypoglycemia. Peak absolute plasma levels of

insulin were similar in both Study 1 (D: 93.2 ± 7.2 ng/mL, D+SSTR2a: 107.9 ± 14.1

ng/mL; Table 4-3) and Study 2 (ExptD2-Ctrl: 71.9±18.7 ng/mL, ExptD2-SSTR2a:

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75.7±12.3ng/mL; Table 5-4), and even exhibited a tendency to be greater in the acute

hypoglycemia experiment of Study 1; thus it is likely that the sensitivity to insulin, rather

than the absolute amount of circulating insulin, is the more important factor for suppressing

glucagon release. We have previously demonstrated that chronic, moderate insulin

treatment administered for the purpose of lessening insulin resistance in STZ-diabetic rats

helps to improve sensitivity to insulin in part, but that these rats still necessitated more

insulin to induce hypoglycemia than non-diabetic rats, despite co-administration of

phloridzin (641).

Another explanation for the diminished capacity of SSTR2 antagonist to fully enhance

hormone responses following recurrent hypoglycemia is that different mechanisms regulate

the defects of counterregulatory hormone responses to acute versus recurrent

hypoglycemia. Thus, while SSTR2 blockade can acutely augment glucagon and

corticosterone in diabetic rats, presumably because elevated pancreatic and/or circulating

somatostatin, it is possible that elevated somatostatin does not contribute to the changes in

neurocircuitry and central mechanisms that are altered with recurrent hypoglycemia. As

discussed in part above, some of these mechanisms include, but are not limited to: (i)

alterations of key brain regions responsible for glucose-sensing (245;660;670); (ii)

alterations of mechanisms responsible for relaying counterregulatory responses

(197;220;641;661;671); and/or (iii) a compensatory increase in glucose and/or other fuel

uptake by the brain (38;178;672). Despite the very modest improvements in hormone

counterregulation attributable to SSTR2 antagonist administration, it is noteworthy to

emphasize that alleviation of the effect somatostatin via SSTR2 resulted in a remarkable

amelioration of hypoglycemia in recurrently hypoglycemic diabetic rats.

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5.5 Study 2 Conclusions

We provide support that recurrent hypoglycemia in diabetic rats markedly attenuates

glucagon and epinephrine responses, as compared to the acute responses in Study 1, to

subsequent hypoglycemia. We demonstrate that hypoglycemia can be ameliorated by

SSTR2 antagonism following recurrent hypoglycemia in diabetic rats by lessening the depth

and duration of hypoglycemia, and this glycemic rescue may be attributable in part to

enhanced glucagon, epinephrine, and/or corticosterone release. We hypothesize that

somatostatin may act via SSTR2 at various sites – pancreas, adrenal, brain, and/or

pituitary, to regulate counterregulation in diabetes but do not exclude other mechanisms of

impairment. Further investigation is necessary to further elucidate the mechanisms by

which euglycemia is restored by inhibiting the action of somatostatin. By minimizing the

risk of hypoglycemia, intensive insulin treatment would be more effective, leading to a lower

incidence of diabetic complications and decreasing the stress associated with the anxiety of

becoming hypoglycemic, consequently improving the quality of life for diabetic patients.

161

Figure 5-1. Study 2 protocol design.

n=13 n=7

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Figure 5-2. Pilot study: A) Glycemia during 2 consecutive days of hypoglycemia (i.e. without prior recurrent hypoglycemia). B) Plasma glucagon levels during 2 consecutive days of hypoglycemia. Total average insulin administered = 4.8 U s.c. Data are represented as mean ± SEM.

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Figure 5-3. Pilot study: A) Glycemia during 2 consecutive days of hypoglycemia (i.e. without prior recurrent hypoglycemia). B) Plasma glucagon levels during 2 consecutive days of hypoglycemia. Insulin administered via i.v. bolus + infusion. Data are represented as mean ± SEM.

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Figure 5-4. Blood glucose levels during recurrent hypoglycemia treatment (5 episodes over 3 days) in rats that will later be divided into SSTR2 antagonist-treatment (saline-SSTR2a; dark grey squares ()) and control (saline-saline; light grey circles ()) groups for hypoglycemia experiments. Data are represented as mean ± SEM.

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Figure 5-5. Blood glucose levels during hypoglycemia experiments in saline-SSTR2a treatment group. ExptD1: black squares (), and ExptD2: white squares (). Data are represented as means ± SEM. An enlarged scale of the period of interest (i.e. euglycemia and hypoglycemia) is presented in Figure 5-7A. †P<0.02 vs. ExptD1

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Figure 5-6. Blood glucose levels during hypoglycemia experiments in saline-saline control group. ExptD1: black circles (), and ExptD2: white circles (). Data are represented as means ± SEM. An enlarged scale of the period of interest (i.e. euglycemia and hypoglycemia) is presented in Figure 5-7B.

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Figure 5-7. Enlarged scale showing blood glucose levels during hypoglycemia experiments in the A) saline-SSTR2a treatment group (solid bars) and B) saline-control group (hashed bars) on ExptD1 (black) and ExptD2 (white). Inset graphs represent extent of hypoglycemia and are calculated as area under the curve of glycemia <4.0 mM. Data are represented as means ± SEM. *P<0.001 vs. ExptD1. †P<0.02 vs. ExptD1

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Figure 5-8. Plasma glucagon during ExptD1 (black) and ExptD2 (white) in A) saline-SSTR2a treatment (solid bars); B) saline-SSTR2a treatment with rats that did not experience hypoglycemia omitted; and C) saline-saline control groups (hashed bars). Data are represented as mean ± SEM. *P<0.04 vs. ExptD1.

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Figure 5-9. Plasma epinephrine during ExptD1 (black) and ExptD2 (white) in A) saline-SSTR2a treatment (solid bars); B) saline-SSTR2a treatment with rats that did not experience hypoglycemia omitted; and C) saline-saline control groups (hashed bars). Data are represented as mean ± SEM. * P<0.03 vs. ExptD1.

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Figure 5-11. Plasma norepinephrine during ExptD1 (black) and ExptD2 (white) in A) saline-SSTR2a treatment (solid bars); B) saline-SSTR2a treatment with rats that did not experience hypoglycemia omitted; and C) saline-saline control groups (hashed bars). Data are represented as mean ± SEM.

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Figure 5-12. Comparison of peak incremental hormone responses to acute hypoglycemia (Study 1; solid bars) and recurrent hypoglycemia (RH) (Study 2; speckled bars) in control (Ctrl; black) and SSTR2 antagonist-treatment (SSTR2a; white) groups. A) Plasma glucagon; B) plasma epinephrine; C) plasma corticosterone. Data are represented as mean ± SEM. *P<0.003 vs. Acute-Ctrl. †P<0.05 vs. RH-Ctrl. ‡P<0.03 vs. Acute-Ctrl.

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Table 5-1. Metabolic parameters for rats that underwent recurrent hypoglycemia treatment. Fed blood glucose, body weight, and food intake were measured daily, and the table shows the mean ± SEM of values averaged over the last week before commencement of recurrent hypoglycemia treatment in rats that would later be divided into SSTR2a-treatment and control groups for hypoglycemia experiments. Daily blood glucose measurements were obtained using a glucometer.

Treatment group

(saline-SSTR2a)

Control group

(saline-saline)

fed blood glucose (mM) 25.3 ± 0.9 23.6 ± 2.1

body weight (g) 365 ± 6 372 ± 11

food intake (g) 39 ± 1 37 ± 2

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Table 5-2. Body weight and amount of insulin administered via iv bolus and iv infusion, and total insulin administered in both groups on both days. Rats received a 10 U/kg iv bolus of insulin after basal samples were obtained at time = 0 min. Subsequently, insulin infusion (50 mU/kg/min) was started and stopped at the experimenter's discretion when hypoglycemia was neared. This volume of insulin infusion was recorded on ExptD1 and repeated on ExptD2. Data are represented as mean ± SEM.

Treatment group

(saline-SSTR2a)

Control group

(saline-saline)

ExptD1 ExptD2 ExptD1 ExptD2

body weight (g) 341 ± 11 340 ± 10 340 ± 11 338 ± 10

initial glycemia (mM) 25.5 ± 1.7 24.1 ± 1.9 20.9 ± 4.0 21.5 ± 4.4

insulin via iv bolus (U) 3.24 ± 0.27 3.26 ± 0.29 3.12 ± 0.63 3.09 ± 0.64

insulin via iv infusion (U) 0.88 ± 0.11 0.90 ± 0.11 0.89 ± 0.30 0.89 ± 0.30

total insulin administered (U) 4.12 ± 0.31 4.16 ± 0.34 4.01 ± 0.86 3.98 ± 0.85

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Table 5-3. Basal plasma hormone levels on Experimental Day 1 (ExptD1) and Experimental Day 2 (ExptD2) in rats that underwent SSTR2a-treated and control hypoglycemia experiments. These represent basal hormone levels after 5 episodes of recurrent hypoglycemia and a partial overnight-fast. Data are represented as mean ± SEM. * P<0.05 vs. ExptD1

Treatment group (saline-SSTR2a) Control group (saline-saline)

ExptD1 ExptD2 ExptD1 ExptD2

glucagon (pg/mL) 58 ± 5 60 ± 6 55 ± 5 53 ± 5

epinephrine (pg/mL) 106 ± 17 102 ± 21 126 ± 33 129 ± 29

corticosterone (ng/mL) 110 ± 20 54 ± 12 * 110 ± 37 132 ± 28

norepinephrine (pg/mL) 326 ± 77 300 ± 77 401 ± 52 378 ± 72

insulin (ng/mL) 0.98 ± 0.2 0.98 ± 0.1 1.00 ± 0.2 0.97 ± 0.2

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Table 5-4. Plasma insulin levels (ng/mL) during ExptD1 and ExptD2 in saline-SSTR2a treatment and saline-saline control groups. Insulin was administered as an i.v. bolus followed by i.v. infusion after samples were obtained at the 0 time point. Infusions were stopped at the experimenter’s discretion (usually between 30-120 min) but kept consistent for the same rat on both experimental days. Data are represented as mean ± SEM.

Plasma insulin (ng/mL)

-60 0 60 180 240

saline-SSTR2a

ExptD1 0.98 ± 0.15 0.90 ± 0.17 96.71 ± 15.94 1.13 ± 0.09 0.70 ± 0.05

ExptD2 0.98 ± 0.15 1.03 ± 0.17 75.71 ± 12.34 1.03 ± 0.07 0.75 ± 0.08

saline-saline

ExptD1 1.00 ± 0.22 1.02 ± 0.22 89.27 ± 32.92 1.43 ± 0.55 0.63 ± 0.14

ExptD2 0.97 ± 0.17 0.93 ± 0.21 71.93 ± 18.73 1.32 ± 0.46 0.60 ± 0.09

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177

Chapter 6 Study Summaries and General Discussion

6 Study Summaries and Limitations

Hypoglycemia remains an obstacle and limitation to stringent glycemic control in diabetes

(4;161). The result of -cell failure in type 1 diabetes and advanced type 2 diabetes causes

the loss of the first two key defenses against a fall in blood glucose levels: the decrement of

insulin and the corresponding -cell response to release glucagon (172). Since glucagon

counterregulation normally plays the primary role in promoting glycemic recovery from

hypoglycemia (510), such that even increases of epinephrine cannot fully compensate for

the absence of glucagon on hepatic glucose production during hypoglycemia (62), it is of

great clinical relevance to discover methods to improve and restore glucagon

counterregulation to hypoglycemia in diabetes. Two main theories underlay the glucagon

deficiency in the present thesis: that in diabetes, the -cell becomes more sensitive to the

inhibitory effects of exogenous insulin, and that excessive somatostatin further inhibits the

ability of the diabetic -cell to release glucagon via SSTR2 agonism during hypoglycemia.

Because somatostatin also has been shown to suppress stress-induced epinephrine and

cortisol/corticosterone release (169;528;628;652;654;656), and because SSTR2 are

present in tissues that store or regulate epinephrine and cortisol/corticosterone secretion

(373;378;379;390;397-400), we also hypothesized that SSTR2 blockade would enhance the

release of these counterregulatory hormones to hypoglycemia. The main objectives of this

thesis were: 1) to evaluate a means to improve the counterregulatory response to

hypoglycemia in a diabetic animal model by use of a SSTR2 antagonist, and 2) to assess if

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counterregulatory hormone improvements could be extended and have a biological effect to

ameliorate hypoglycemia after exposure to recurrent hypoglycemia.

6.1 Summary of Study 1

Data from the present study demonstrate that diabetic rats had markedly attenuated

glucagon and corticosterone responses to hypoglycemia as compared to non-diabetic rats,

which is in agreement with our previous studies (205). We demonstrate that glucagon and

corticosterone counterregulatory hormone responses to hypoglycemia can be restored in

diabetic rats with specific antagonism of the SSTR2 using both high (10 U/kg) and low (5

U/kg) doses of insulin to induce hypoglycemia. We attribute these defects in part to

elevated pancreatic and circulating somatostatin levels which were increased in diabetic rats

following hypoglycemia. SSTR2 antagonist given to non-diabetic rats, in which somatostatin

levels were the same as non-diabetic controls, did not increase glucagon or corticosterone

responses, which further provides support for the contribution of augmented somatostatin.

During basal conditions in the absence of insulin, this SSTR2 antagonist does not elicit

hyperglycemia or substantial elevations in counterregulatory hormones, which is important

if this antagonist is potentially considered for therapeutic use in humans. Under conditions

of lower dose insulin injection, SSTR2 antagonism reduced the glucose requirement during

hypoglycemic clamp, which is consistent with improved glucagon and corticosterone

counterregulation and is indicative of a lower risk of hypoglycemia. In these 3-week STZ-

diabetic rats, epinephrine and norepinephrine counterregulation to hypoglycemia induced

with both high and low doses of insulin were similar in both non-diabetic and diabetic rats;

thus, these rats did not demonstrate a defect in catecholamine responses to hypoglycemia.

Epinephrine, norepinephrine, and growth hormone levels were not affected by SSTR2

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administration. Despite improvements of glucagon and corticosterone to hypoglycemia,

expression of gluconeogenic enzymes PEPCK1 and G6Pase were unaltered.

6.2 Summary of Study 2

In light of the evidence from Study 1 that SSTR2 antagonism could normalize glucagon and

corticosterone response to hypoglycemia, we designed a set of experiments to determine

whether such improvements in hormone responses could have a biological effect and

improve recovery from hypoglycemia in a model with established counterregulatory defects:

recurrent hypoglycemia. In these experiments, we first subjected diabetic rats to 5

episodes of hypoglycemia. Subsequently, we compared the responses of each rat over two

experimental days. Much care was taken to control for similar baseline metabolic

parameters, such as initial glycemia, body weight, and overnight food intake, so that

meaningful and relevant comparisons between the two experimental days could be made.

On the first experimental day (ExptD1), we determined the minimum amount of insulin

necessary to induce hypoglycemia at 3.0 mM in a given rat, which was necessary since

diabetic rats have varying insulin sensitivities. On the second experimental day (ExptD2),

we used to the same dose of insulin in the same rat and compared its glycemic and

hormone responses to ExptD1. We included two groups in this study: a treatment group

which received SSTR2 antagonist on ExptD2, and a control group which did not receive

SSTR2 antagonist on ExptD2. By comparison with Study 1, we demonstrated that recurrent

hypoglycemia further reduced the attenuated glucagon response, and markedly reduced the

epinephrine response, to subsequent hypoglycemia in diabetic rats. Our most striking

finding was that hypoglycemia can be ameliorated by SSTR2 antagonism in diabetic rats,

despite the marked attenuation of glucagon and epinephrine caused by recurrent

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hypoglycemia. By blocking the actions of somatostatin on SSTR2, the depth and duration of

hypoglycemia were significantly lessened, and a quicker recovery to euglycemia ensued in

animals, despite prior exposure to repeated hypoglycemia. This amelioration of

hypoglycemia may be due, at least in part, to enhanced glucagon release and to

improvements in epinephrine and corticosterone counterregulation which are observed in

the SSTR2 antagonist treatment group. Norepinephrine levels were not affected by

recurrent hypoglycemia or by SSTR2 antagonist treatment.

6.3 General discussion

Our results suggest an important role for increased pancreatic, and possibly circulating,

somatostatin in defective glucagon, corticosterone, and/or epinephrine counterregulation in

diabetes but do not exclude other mechanisms of impairment. From our present findings, it

is hypothesized that selective SSTR2 blockade at the levels of the brain, pituitary, pancreas,

and adrenal gland during hypoglycemia may help to reduce the severity, duration, or onset

of hypoglycemia by enhancing the release of counterregulatory hormones such as glucagon,

corticosterone, and epinephrine.

6.3.1 Limitations of the present study

The main limitation is that a precise mechanism of action of the SSTR2 antagonist could not

be delineated from the work of this thesis. From the data presented in this thesis, we can

suggest that SSTR2 antagonism plays a role in restoring glucagon and corticosterone

counterregulation to acute hypoglycemia, and has some effect to improve glucagon,

epinephrine, and corticosterone responses following recurrent hypoglycemia, but we have

yet to delineate the precise mechanism by which this occurs. These impairments in

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hormone responses can occur at the tissue level (pancreatic islets for glucagon, adrenal

cortex for corticosterone, or adrenal medulla for epinephrine), via the peripheral nerves, or

at the level of the brain or pituitary. We suggest that increased pancreatic somatostatin,

and potentially increased circulating somatostatin, can affect local and central control of

counterregulatory hormone release. With respect to deficient glucagon counterregulation,

we hypothesize that despite the fact that somatostatin may not be excessive in certain

models of diabetes, antagonism of somatostatin’s suppressive actions may still be beneficial

since diabetic -cells are already more sensitive to the inhibition of insulin. With respect to

impaired corticosterone counterregulation, we hypothesize that an additional alleviation of

suppression of corticosterone release may improve this response to hypoglycemia. Our

laboratory has previously demonstrated that in diabetes, corticosterone release to

hypoglycemia is attenuated by suppression on the HPA axis (202;220;641;673). Although

we did not delineate a role for somatostatin on this inhibition, we presently hypothesize that

somatostatin may inhibit activation of the HPA axis since somatostatin is known to inhibit

release of neurotransmitters and hormones involved in the HPA response, namely

norepinephrine, CRH, and ACTH. However, the limitation remains that studies to assess the

direct effect of SSTR2 blockade in the hypothalamic PVN (which could be achieved by

stereotaxic microinjection of the SSTR2 antagonist) or in corticotropes of the anterior

pituitary were not performed. Measurement of plasma ACTH could also provide an

indication as to whether inhibition of somatostatin involved an effect at the level of the

anterior pituitary. The anterior pituitary is not protected by the blood-brain-barrier, and

peripheral administration of the SSTR2 antagonist could theoretically bind with SSTR2 in the

anterior pituitary. Thus, if the SSTR2-mediated improvements in corticosterone

counterregulation are accompanied by increased ACTH levels, this would indicate a direct

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effect of disinhibition at the level of the corticotrope or suggest an involvement of a central

mechanism.

In addition, another limitation is that we did not measure glucose turnover in these studies.

Precise and accurate measurements of glucose turnover could have helped us to determine

whether the enhancement of counterregulatory hormones and improvement in

hypoglycemic recovery associated with SSTR2 blockade had an effect on glucose

production, glucose utilization, or both. Further investigation is therefore necessary to

elucidate the mechanism by which hormone counterregulation is improved and euglycemia

is restored via inhibiting the action of somatostatin via SSTR2.

6.3.2 Hypothesizing a mechanism of action: CNS effects?

Our initial hypothesis was that the concomitant elevation of pancreatic somatostatin and

heightened sensitivity of diabetic -cells to exogenous insulin contributed to the impairment

of the glucagon response to hypoglycemia in diabetes. This, of course, cannot explain the

improvements of corticosterone and epinephrine counterregulation with SSTR2 disinhibition.

In the Discussions of Study 1 and Study 2, several potential mechanisms were addressed

that may underlie SSTR2-mediated improvements to glucose counterregulation. In addition

to these, this section will briefly propose possible roles of SSTR2 antagonism within the

brain that could regulate counterregulatory hormone responses.

There is vast, compelling evidence dating back to early studies of Claude Bernard in the

19th century in which he demonstrated that punctures of the fourth cerebral ventricle

rendered dogs glycosuric (674). Recent reviews (13;14;193) on brain glucosensing and

glucose counterregulation also irrevocably demonstrate the importance of the central

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nervous system on regulating glucose homeostasis. Initiation of counterregulatory hormone

responses in the central nervous system involves: (i) afferents that detect changes in

glucose levels in the brain and/or periphery; (ii) integration and relay of signals within

specific brain regions; and (iii) efferent pathways from integration centres in the brain that

project as autonomic signals to the body to maintain glucose homeostasis (13;14;193).

Interestingly, many of these brain regions that comprise the neural networks that regulate

hypoglycemic counterregulation also are rich in somatostatin and its receptors. This is not

surprising since within the brain, somatostatin serves as a neurotransmitter. Yet it allows

us to speculate: could alterations in brain somatostatin action as a result of diabetes and/or

recurrent hypoglycemia contribute to the impairments of hypoglycemic counterregulation?

In addition to mechanisms within the pancreas responsible for glucagon release in response

to hypoglycemia (76), glucagon counterregulation is also regulated centrally. Buijs et al.

used a pseudorabies virus as a retrograde transsynaptic tracer to elucidate the neuronal

parasympathetic and sympathetic efferent pathways that control pancreatic hormone

secretion (99). Buijs et al. identified a sympathetic pathways involving (i) neurons of the

ventromedial hypothalamus (VMH), arcuate nucleus (ARC), central amygdala, bed nucleus

of the stria terminalis (BNST), suprachiasmatic nucleus (SCN), and medial preoptic area

(MPO) projecting to the paraventricular nucleus (PVN) and hypothalamic zona incerta (ZI),

which in conjunction with a projection from the lateral hypothalamus (LHA) project to the

intermediolateral column (IML) of the spinal cord; and (ii) neurons of the nucleus of the

solitary tract (NTS) projecting to the locus coeruleus (LC) of the brain stem, which

subsequently joins a projection from the prefrontal cortex, or from the NTS to the

rostroventrolateral medulla (RVLM) to project to the IML. Both of these neuronal pathways

subsequently projecting from the IML of the spinal cord provides the sympathetic efferent to

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the pancreas to stimulate glucagon release (99). In addition, in a recent review, Watts and

Donovan summarized the interactions between networks involving the periphery, hindbrain,

and hypothalamic components involved with hypoglycemic counterregulation (14). In brief,

glucose sensors are located in the portal/mesenteric vein (PMV), hindbrain, and

hypothalamic nuclei (LHA, ARC, and VMH). These signals are then integrated in networks

involving the hindbrain and hypothalamus. The hindbrain and hypothalamic integrators

then activate autonomic efferent neurons to stimulate glucagon and catecholamine release,

or via a neuroendocrine route the PVN stimulates glucocorticoid release (14). In his review,

Cryer has also indicated key brain regions that participate in the mechanisms of

hypoglycemia-associated autonomic failure, which include the prefrontal cortex, the dorsal

midline thalamus, hypothalamus, hippocampus, brain stem, and amygdala (4).

These recent reviews identify key brain areas involved with hypoglycemic counterregulation.

Within the brain, somatostatin-immunoreactivity is found in many neurons. As reviewed by

Viollet et al., somatostatinergic neurons are highly detected in the mediobasal

hypothalamus, median eminence, amygdala, hippocampus, cerebral cortex, and brainstem

(292). In addition to short projections that act within a local area, somatostatinergic

neurons can also project to other target areas, and several examples include: amygdala

parabrachial nucleus (PBN), NTS, and dorsal motor nucleus of the vagus (DMNV);

periventricular nucleus median eminence, ARC, habenula, and LC; PBN thalamus; and

NTS PVN, PBN. SSTR2 expression has been reported in hypothalamic ARC, PVN, and

VMH (420;424;426), medial habenular nucleus of the thalamus (420), cerebral cortex,

hippocampus, LC, basolateral amygdala (BLA), NTS, PBN, and spinal cord (419;420;425).

In light of the fact that several SSTR2-expressing brain regions endogenously are targeted

by somatostatinergic neurons, and that these regions also play a role in glucose-detection

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and/or regulation of the counterregulatory responses, it would be of interest to evaluate the

effect of specific SSTR2 antagonism on glucose-regulatory neuronal populations. This could

be achieved by inhibiting these receptors during hypoglycemia by directly administering the

SSTR2 antagonist to specific brain regions or by silencing the SSTR2 gene at specific brain

nuclei via viral gene delivery. Also, it would be interesting to determine whether these

glucose-regulatory neurons are affected by diabetes and/or hypoglycemia. This could

provide new insights for the role of somatostatin in diabetic hypoglycemic counterregulation.

6.3.2.1 Does peripherally administered SSTR2 antagonist enter the brain?

We are not aware at the present time of any reports whether our SSTR2 antagonist can

enter the brain when administered peripherally via an intravenous route. CTAP, a cyclic

octapeptide with the structure D-Phe-Cys-Tyr-D-Trp-Arg-Thr-Pen-Thr-NH2 is a -opiod

receptor antagonist that has been reported to cross the blood-brain-barrier (562).

However, the majority of literature, however, suggests that a compound like our cyclized

octapeptide antagonist (see Figure 1-2 for structure) would likely not cross the blood-brain-

barrier (675). Octreotide, an octapeptide somatostatin analogue, is unlikely to cross the

blood-brain-barrier in any significant amounts when peripherally administered (676).

Furthermore, SSTR3 octapeptide antagonists cyclized via a disulfide bridge have been

reported to be very poor at crossing the blood-brain-barrier (677). In view of these reports,

our peripherally infused SSTR2 antagonist may not have entered the brain to have any

central effects. Further investigation is needed to more precisely determine whether this

compound could enter the brain. The MDCK-MDR1 cell line is an established in vitro model

of the blood-brain-barrier and can be used to test the permeation properties of peptides

(678).

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6.3.3 Brief evaluation of tachyphylactic effects of SSTR2 antagonism

It is known that G-protein coupled receptors (GPCRs), including SSTRs, share a common

property whereby they regulate their own responsiveness to continued agonist exposure

(679). For example, GPCRs become desensitized after acute treatment to agonists. Three

main steps are responsible for this: 1) uncoupling of the GPCR from G proteins; 2) receptor

internalization via the phosphorylation of the C-terminal tail and intracellular loops by

second-messenger-activation or GPCR-kinases, which has been demonstrated in rat

pituitary and islet tumour cells; and 3) receptor downregulation. SSTR2, for example

couples with adenylyl cyclase via inhibitory G proteins when activated by somatostatin and

undergoes endocytosis via clathrin-coated pits (680). However, GPCRs also become

upregulated after prolonged agonist exposure (679). This is associated with: 1) the active

process of ligand-induced receptor recruitment from a preexisting cytoplasmic pool; 2) does

not require new protein synthesis; and 3) depends on molecular signals on the C-terminal

tail. This has also been observed in receptors for somatostatin (681). These findings are

relevant for agonist exposure. However, it remains to be seen if the same applies for

antagonist exposure. For our SSTR2 antagonist used in this study, in which the peptide

ligand binds to the SSTR2 without inducing the same intracellular responses as somatostatin

would, it is not known how prolonged exposure would affect receptor expression on target

tissues, such as pancreatic -cells, the brain, or adrenal glands. There is no consensus in

literature as to whether antagonists initiate internalization of GPCRs. Peptide receptor

antagonists for cholecystokinin (682) and neuropeptide Y (683), for example, have been

reported to stimulate receptor internalization. One study demonstrated that SSTR agonists,

but not antagonists internalized SSTR2 (680). Another study likewise demonstrated that

antagonists for SSTR2 or SSTR3 did not affect internalization in human embryonic kidney

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cells (684), but studies for our specific antagonist and its result in -cells are still required

to elucidate the long-term efficacy of this peptide.

6.3.4 Potential side effects of SSTR2 antagonism

Since receptors for somatostatin are expressed ubiquitously throughout the body, it is

important to target the particular tissue – and specifically the cell type – in which a

pharmacological compound is to take effect. In particular, we initially sought to target

SSTR2 because of its mediation of somatostatin’s inhibitory effect on -cell glucagon

release. Using an antagonist with high selectivity, binding affinity, and biological activity is

of key importance, and the peptide antagonist that was used in this study offered these

characteristics (509). However, although most tissues tend to show preferential expression

for particular SSTR subtypes, SSTR2 is still expressed in high levels in several areas. These

areas include the stomach, the cerebral cortex, pancreatic -cell, and adrenal medulla. In

the pituitary, SSTR5 has the highest level of expression, followed by SSTR2. The high

expression of SSTR2 on -cells and the adrenal medulla is of course favourable for our

studies in glucagon and epinephrine counterregulation. These other tissues/cell types are

mentioned here to assess potential side effects of SSTR2 antagonist administration during

our experiments.

Although pancreatic -cells in rodents exclusively express SSTR5 and not SSTR2 (387), high

levels of SSTR2 antagonist without sufficient competition by native somatostatin or with

saturation of SSTR2 on -cells may lead to non-specific binding with SSTR5. This would

lead to enhanced insulin secretion from functional -cells, including those in non-diabetic

subjects or possibly in non-advanced type 2 diabetes. This may explain why diminished

glucagon responses to hypoglycemia were observed in our studies of SSTR2 antagonist in

non-diabetic rats. This would not be a concern, however, in type 1 diabetes or likely not in

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STZ-diabetes where the majority of -cells are destroyed. Furthermore, although human -

cells predominantly express SSTR1 and SSTR5, SSTR2 is expressed in up to 25% of human

-cells, which may suggest a greater potential side effect of SSTR2 antagonist in promoting

insulin release. Thus, patients with type 2 diabetes who still have insulin secretory capacity

should avoid SSTR2 antagonist as a therapy to prevent hypoglycemia.

In the stomach, SSTR2 has been reported as the main subtype through which somatostatin

inhibits gastrin-stimulated gastric acid secretion both through studies using SSTR2 peptide

analogs (685) and SSTR2 knockout mice (686). A study using the same SSTR2 antagonist

in this present work demonstrated in non-diabetic rats that SSTR2 antagonism blocked the

inhibitory effect of somatostatin on gastrin-stimulated release. At a dose which we used in

our pilot studies (1500 nmol/kg/h), the SSTR2 antagonist likewise inhibited the suppressive

effect of GLP-1 on gastrin-stimulated gastric acid output (508). However, a recent study

reported normal circulating levels of gastric and unaltered gastric acid output in SSTR2

knockout mice (687), which suggests that blocking the effects mediated via SSTR2 may not

necessarily affect gastric function. However, it remains possible that in our studies,

stimulated-gastric acid output could have been increased in our rats. Other gut effects

mediated by somatostatin action via SSTR2, such as suppression of intestinal muscosal cell

proliferation and colonic motility, may also be inhibited by SSTR2 antagonism. Although

glucose absorption was shown to be inhibited by somatostatin (688), the receptor subtype

by which this action is mediated was not determined.

In the brain, somatostatin functions as a neurotransmitter contributing to cognitive,

locomotor, sensory, and autonomic activities (271). SSTR1 is considered by many reviews

to be more vastly distributed and highly expressed than other receptor subtypes, and is

often found in higher levels than SSTR2 in areas such as the cerebral cortex and

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hippocampus where SSTR2 is considered to be highly expressed (271;378;420).

Somatostatin was initially known for its effect to suppress growth hormone secretion (267).

Somatostatin produced in the periventricular nucleus (where hypothalamic somatostatin is

greatest (271)) and released from the median eminence directly inhibits growth hormone

release from somatotrophes of the anterior pituitary via SSTR2 and SSTR5 (292;431).

SSTR5 is the main subtype expressed throughout the pituitary, followed by SSTR2 (380).

There is also evidence that SSTR1 is also implicated in the control of growth hormone

release (433). Whereas SSTR2 antagonist disinhibition to stimulate growth hormone

secretion would be favourable during hypoglycemia, this was not observed in our present

findings. However, it remains to be determined whether prolonged treatment with SSTR2

antagonist, or blood sampling at more frequent intervals, could stimulate growth hormone

release. Presently, we have no direct data to report whether SSTR2 antagonist stimulated

anterior pituitary secretion of adrenocorticotrophic hormone (ACTH), but our improved

corticosterone data with SSTR2 antagonist treatment could support this. Other studies have

demonstrated an inhibitory effect of somatostatin on ACTH release (294) and the presence

of SSTR2 on ACTH cells in the anterior pituitary (561). Other pituitary hormones, such as

thyroid stimulating hormone and adrenocorticotrophic hormone, may also be altered by

SSTR2 antagonism (290).

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Chapter 7 Future Directions

7 Future Directions

7.1 Delineating a mechanism of action for SSTR2 antagonism

To address the main shortcoming of this project, which is a lack of mechanistic explanation

for the SSTR2-antagonism mediated improvements in glucose counterregulation, an

important future study would be to delineate a mechanism by which SSTR2 blockade acts

to: (i) improve hormone counterregulation to acute hypoglycemia; (ii) more effectively

restore low blood glucose back to euglycemia. For the latter, tracer experiments using

traditional hyperinsulinemic-hypoglycemic clamp techniques can provide glucose turnover

data which may provide insights as to whether somatostatin affects glucose production or

uptake during hypoglycemia. This is important since we have previously shown that peak

glucose production was decreased during hypoglycemia in diabetic rats (205). An

improvement in net glucose production is essential since the overall purpose in improving

counterregulation is to restore plasma glucose levels in diabetic patients suffering from

hypoglycemia. However, despite the unspecified etiology of the effect, a significant

improvement in glycemic recovery was demonstrated by SSTR2 inhibition, and it is often

the overall effect of a pharmacological compound that garners the interest and attention of

patients.

To determine whether normalized glucagon and corticosterone counterregulation was

mediated via a central nervous system effect, SSTR2 antagonist can be

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intracerebroventricularly administrated during hypoglycemia in diabetic rats. Finally,

evaluating the counterregulatory hormone responses to hypoglycemia in genetic animal

models with tissue-specific SSTR2 knockout can also help to elucidate mechanisms of

somatostatin action.

7.2 Testing the efficacy of SSTR2 antagonism in other diabetic models

In light of the fact that certain diabetic rodent models, such as the BB rat, which do not

have increased pancreatic somatostatin, but do have elevated circulating somatostatin

(537;689) do show impairments in the counterregulatory hormone responses to insulin-

induced hypoglycemia (263;542;543), it is worthwhile to investigate the efficacy of SSTR2

antagonism in this model of experimental diabetes to determine the contribution of

excessive pancreatic somatostatin on glucagon counterregulation in diabetes. It is unknown

why BB rats do not have increased pancreatic somatostatin, unlike other experimental rat

models of diabetes. An earlier hypothesis suggested that BB rats had “more severe islet

destruction” than STZ-diabetic rats (537). In addition, it has been demonstrated that

insulin treatment of STZ-diabetic rats can normalize, at least in part, pancreatic and

circulating somatostatin levels, yet insulin-treated diabetic animals do not have fully

normalized hormone counterregulation to hypoglycemia (174;202;641). Thus, performing

similar hypoglycemia studies with and without SSTR2 antagonist in insulin-treated rats will

allow us to determine whether improvements in diabetic hyperglycemia and -cell

sensitivity may alter the effectiveness of SSTR2 antagonism during hypoglycemia. Insulin-

treated diabetic subjects may also be less insulin resistant, which would allow us to use less

insulin to induce hypoglycemia. This is important since excessively high insulin levels alter

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counterregulation and glucose production. Insulin-treated diabetes is more clinically

relevant since type 1 diabetic humans are insulin treated.

7.3 Effect of SSTR2 antagonism to non-hypoglycemia stimuli

It is presently unknown whether SSTR2 antagonism will enhance glucagon release in

response to other stimuli known to trigger glucagon release, such as an amino acid-rich

meal or exercise. Hypoglycemia and amino acids stimulate glucagon secretion by two

different processes, the latter of which only can be counteracted by amylin (690). Exercise

and hypoglycemia similarly stimulate glucagon via activation of the autonomic nervous

system and neuroendocrine response (149). Unlike hypoglycemia, the glucagon response

to amino-acid rich meals (691-694) and exercise (149;695-697) are preserved, or even

exaggerated, in diabetes. Thus, studies designed to evaluate the effects of SSTR2

antagonist on other glucagon secretagogues will be of interest, and it may help to

determine if this effect to enhance counterregulatory hormones is hypoglycemia-specific.

Plasma glucagon could be measured in response to: i) a high protein meal, and ii) exercise

with and without administration of the antagonist, in the presence or absence of elevated

somatostatin, and in non-diabetic or diabetic subjects, to determine the stimulus-specificity

of SSTR2 inhibition and the condition(s) in which it applies.

7.4 Interaction between glycemia, insulin, and somatostatin antagonism on glucagon release

Using perfused pancreata or isolated islets, the direct effect of the SSTR2 antagonist on

glucagon release can be assessed with carefully controlled interactions between glycemia,

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insulin, and somatostatin antagonism on glucagon release. These ex vivo or in vitro

experiments have the benefit of clarifying the role of somatostatin antagonism in the

pancreas more directly since in vivo experiments also involve many other hormonal and

metabolic changes. Efendic et al. has shown previously that this SSTR2 antagonist can

increase arginine-induced glucagon release in perfused pancreata at both 5.5 and 3.3 mM

glucose concentrations in non-diabetic rats (392). However, those experiments did not

answer the main question of whether SSTR2 antagonist can, in presence of insulin which

inhibits -cells, increase glucagon release in diabetic islets, which are often deprived of the

tonic effect of insulin. These experiments can also assess the contribution of increased -

cell sensitivity to insulin as a result of a chronic lack of the presence of insulin on glucagon

secretion to insulin-induced hypoglycemia. These experiments could also show whether the

local effect on glucagon release is as marked as when SSTR2 antagonist is given

peripherally.

194

194

Chapter 8 Conclusions

8 Conclusions

8.1 Importance of contributions to current body of knowledge

Our laboratory first suggested 20 years ago that increased somatostatin in diabetes may

contribute to the impaired glucagon response to hypoglycemia in diabetes (90). We

hypothesized that the combined suppressive effects of excessive pancreatic somatostatin

and increased sensitivity of diabetic -cells to exogenously administered insulin contributes

to defective glucagon counterregulation. The work in this thesis demonstrates for the first

time that antagonizing the actions of somatostatin acting on SSTR2 can fully abolish this

glucagon defect in diabetes during acute hypoglycemia. Thus, disinhibition of the diabetic

-cell from somatostatin, one of its key suppressing influences, can improve its secretory

capacity in response to hypoglycemic stimulation. We also demonstrate that the attenuated

corticosterone response in diabetic rats was significantly improved with SSTR2 blockade, an

effect which at the present time can only be speculated to due to alleviation of

somatostatin’s inhibitory release of corticosterone, or other regulators of its release directly

at the adrenal cortex or indirectly mediated in the brain or pituitary in diabetes. These

improvements of glucagon and corticosterone were observed to be diabetes-specific since

SSTR2 antagonism in non-diabetic rats during hypoglycemia did not increase these hormone

responses. Importantly, we also demonstrate that during basal conditions in the absence of

insulin that SSTR2 antagonist treatment neither causes marked sustained elevations of

counterregulatory hormones nor elicits undesired hyperglycemia in diabetic rats. In

195

addition, we provide novel evidence that SSTR2 antagonism can promote the recovery of

hypoglycemia, presumably in part by improving glucagon, and preserving epinephrine and

corticosterone, hormone responses in diabetic rats exposed to repeated episodes of

hypoglycemia. This exciting and novel finding reveals that the depth and duration of

hypoglycemia can be significantly be reduced and even prevented, even in diabetic subjects

with marked counterregulatory defects. Taken together, these findings implicate an

important role for increased pancreatic, and possibly circulating, somatostatin in defective

hypoglycemic counterregulation in diabetes and do not discount other mechanisms of

impairment, which have been discussed in this thesis.

8.2 Potential clinical contributions

The goal of this work was to develop a means to ameliorate hypoglycemia. This objective

differs from the current hypoglycemia therapies, such as injectable glucagon and dextrose,

which are reactionary measures as opposed to preventive therapies. Hypoglycemia that is

particularly dangerous includes that which occurs undetected, such as during sleep and in

diabetic children and adolescents. Reactionary therapies necessitate the recognition of

hypoglycemia by the individual or the care giver, which would be difficult with hypoglycemic

unawareness or during nocturnal hypoglycemia. A prophylactic solution involves

administration of a therapeutic prior to hypoglycemia onset so that the duration/severity of

hypoglycemia is lessened or prevented entirely so that the patient does not become

debilitated. This could involve administration of the SSTR2 antagonist with one’s insulin,

particularly when the risk of hypoglycemia may be greater (i.e. at bedtime and/or after

exercise). By minimizing the risk of hypoglycemia, intensive insulin treatment would be

more effective, leading to a lower incidence of diabetic complications and decreasing the

196

stress associated with the anxiety of becoming hypoglycemic, consequently improving the

quality of life for diabetic patients.

197

197

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