appetite and adiposity of the emu (dromaius novaehollandiae)

Appetite and adiposity of the emu

(Dromaius novaehollandiae).

by

Gemma Kate Graham

B.Sc. (Agric, Hons)

This thesis is presented for the degree of

Doctor of Philosophy of Animal Science

2002

The University of Western Australia

Faculty of Agriculture

Animal Science Group

Dedicated to

Jean Eiffler and Jim Graham

Who helped me choose a road less travelled.

Declaration

This thesis contains no experimental material that has been previously presented for any

degree at any university or institution. The experimental designs and manuscript

preparation were performed by myself, in consultation with m y supervisors, Dr Dominique

Blache, Dr Philip Vercoe and Professor Graeme Martin.

G e m m a Graham

March 2002

General Index

(Detailed indices are provided at the start of each chapter)

Page

Summary ii

Acknowledgements iv

Publications vi

Index to figures and tables vii

General Introduction 1

Chapter 1 Literature Review 3

Chapter 2 General Materials and Methods 29

Chapter 3 The emu obese gene 58

Chapter 4 The influence of dietary fat on food intake and adiposity 79

Chapter 5 The effect of dexamethasone on food intake, adiposity, 99

reproduction and the hypothalamic expression of N P Y in females

Chapter 6 Incubation-related changes in food intake, adiposity and the 125

hypothalamic expression of N P Y and VIP

Chapter 7 The effect of short-term starvation on the hypothalamic 146

expression of N P Y and VIP during a period of elevated appetite

General Discussion 162

Bibliography 166

Appendix 1. - abbreviations 202

Appendix 2. - list of suppliers and feed composition 205

Appendix 3. - response to comments made by thesis examiners 209

ii

Summary

Emus exhibit seasonal decreases in appetite and adiposity. Adipose tissue is the most

profitable component of the carcass. As such, the reduction in adipose stores limits

slaughtering to a brief window, and is a major constraint to productivity and industry

development. To minimise the detrimental effects of seasonality on the productivity of

farmed emus it is necessary to develop ways of manipulating appetite and adiposity. The

general model for the control of appetite and adiposity is mediation of appetite and satiety

centres in the hypothalamus by peripheral signals indicative of energy reserves and

metabolic status. The relevance of this general model to the emu has not been

investigated.

The general hypothesis in this thesis was that seasonal variation in appetite and adiposity

in the emu operates under the general model developed in other species. The experimental

models used to test the general hypothesis were increased appetite and adiposity (high fat

diet, dexamethasone treatment), decreased appetite and adiposity (incubation) and

increased appetite and decreased adiposity (starvation). To determine the role of leptin in

the control of appetite and adiposity attempts were made to clone and sequence the emu

leptin gene. The concentrations of metabolic hormones were measured in all of the

experiments, and the role of N P Y in the control of appetite and adiposity was investigated

by quantifying the m R N A expression of N P Y from the mediobasal hypothalamus and

preoptic area in the glucocorticoid treatment, incubation and starvation experiments.

NPY mRNA expression did not change in response to dexamethasone treatment or

starvation. There was no difference in N P Y expression between incubating and non-

incubating emus during the breeding season, despite large differences in their appetite and

adiposity. Attempts to clone and sequence emu leptin were unsuccessful and further

investigation of the relationship between leptin and N P Y was not possible. However, it

was demonstrated that emus have biologically active leptin by using a chicken leptin

bioassay developed in Friedman-Einat's laboratory. High fat diets and dexamethasone

treatment did not increase adiposity or food intake. Emus were relatively tolerant of

starvation. Short-term starvation did not increase corticosterone concentration or N P Y

expression, or induce the classical starvation response in metabolic hormone

concentrations that, in other species, would normally occur rapidly in response to

starvation. The hormone concentrations measured throughout these experiments were

appropriate to the physiological requirements of the birds, and indicate that efficient

mechanisms exist to defend the "set point" for energy reserves of the e m u at a level

appropriate to its needs and activities.

The fact that NPY gene expression is unchanged by starvation, incubation or

dexamethasone treatment does not rule out a role for N P Y in the hypothalamic control of

appetite. Other appetite mediators such as, leptin or corticotrophin-releasing hormone

(CRH), could, under the influence of photoperiod, inhibit N P Y action or responses.

Finally, it is apparent from the results that appetite and adiposity are tightly regulated in

the emu. The e m u also shows greater resistance to manipulation and tolerance of

starvation than other species. These features make disrupting energy balance to increase

adiposity a difficult enterprise for the farmer and, for the present, not feasible.

iv

Acknowledgements

I would like to offer m y sincere thanks to the following people:

• My supervisors, Dr Dominique Blache, Dr Philip Vercoe and Professor Graeme

Martin, for their support, guidance and friendship.

• The academic staff of the Animal Science Group at the University of Western

Australia, Emeritus Professor David Lindsay, Emeritus Professor Reg Moir, Professor

Graeme Martin, Dr Ian Williams, Dr John Milton, Dr Philip Vercoe, Dr Dominique

Blache, Dr Suzannah Williams, Dr Irek Malecki and Dr Roberta Bencini for their help

and friendship throughout m y endeavours.

• Technical and support staff past and present in the department, John Beesley, Rob

Davidson, Margaret Blackberry, Margaret Lalchere, Jen Slater, John Koppen and

Sarah-Jo Smith for lending a helping hand, supporting m y efforts and their valued

friendship.

• I would especially like to thank John Beesley for his ability to find what was needed

and somehow the money to pay for it, and Margaret Blackberry for helping in every

aspect of m y research. Without both of you this would never have been completed.

• My fellow postgraduate students completed and continuing, Lucia Chagas, Pietro Celi,

Agung Riono, Tejinder Sharma, Rachel Smith, Matt Thyer, Megan Trezona, Harriet

Mills, Sid Saxby, Dean Thomas, Kristen Wolfe, Craig Lawrence, Justin Bellanger,

Ahmed Ali, Judy Van Cleeff, Karen Williams, U m a Karki, and Georgett Banchero, for

their friendship, advice, invaluable assistance, support, counselling and laughter.

• The Rural Industries Research and Development Corporation for providing my

scholarship and assisting in m y travels as part of this research.

• Leon and Ray in the workshop, for all their fix-its and help.

V

• Peter Cowl at Shenton Park for his help with the emus and for taking such excellent

care of them.

• At Curtin University, Dr Emma Hammond for generously providing her emu cosmid

library and technical advice.

• Dr Christopher Ashwell, USDA, for the gift of vector DNA containing part of the

chicken leptin gene.

• At CSIRO, Dr Norm Adams and Mrs Jan Briegel for advice and encouragement.

• At the Roslin Institute, Scotland, Professor Peter Sharp and Dr Tim Boswell for their

support, advice and encouragement, and Beth Baines for her wonderful hospitality.

• At the Volcani Center, Israel, Dr Miri Einat and family for making my visit both

productive and enjoyable.

• Tom Trevlett from Davison Industries for the kind donation of full-fat canola meal for

m y first experiment.

• My parents, Heather and Laurie, for encouraging me to do whatever made me happy

and whose ability to overcome their dragons has always inspired me, and made m e

very proud.

• And last but by no means least, Daniel, who helped out with my experiments, rejoiced

in m y achievements, held m y hand and stopped m e from taking myself too seriously.

VI

Publications

Work presented at scientific meetings:

Graham G.K., Blache D., Blackberry M., Martin G.B.M. and Vercoe P.E. (1999) Partial

cloning of the emu obese gene sequence. Proceedings of the Combio 99

conference.

vii

Index to figures and tables

Figures

Page

1.1: Annual changes in feed intake and live weight of male and 5 female emus.

1.2: The control of food intake and adipose tissue deposition. 11

1.3: The hypothalamic-pituitary-adrenal axis: effects of endogenous 24 and exogenous glucocorticoids.

2.1: Live weight measurement of the emu. 31

2.2: Jugular bleeding of the emu. 33

2.3: An emu skull showing the placement of incisions through the 35 skull for brain removal.

2.4: Schematic saggital view of the avian brain showing the 35 microdissection sites for collection of the mediobasal hypothalamus ( M B H ) and preoptic area (POA) for R N A extraction and RPA.

4.1: Average daily feed intake for each week of the experiment for 85 male emus receiving either the low or high fat diet for 8 weeks.

4.2: Mean live weight of male emus offered low and high fat diets for 86

54 days.

4.3: Serum glucagon concentrations of male emus offered low and 88 high fat diets for 54 days.

4.4: Serum insulin concentration of male emus offered low and 89 high fat diets for 54 days.

4.5: Serum insulin/glucagon ratio of male emus offered low and 90 high fat diets for 54 days.

4.6: Serum triiodothyronine concentration in male emus offered low 91

and high fat diets for 54 days.

4.7: Serum thyroxine concentration in male emus offered low and 92

high fat diets for 54 days.

4.8: Serum T3/T4 ratio in male emus offered low and high fat diets for 93

54 days.

Figures (continued)

Page

5.1: The serum corticosterone concentration of emus treated with 105 subcutaneous dexamethasone implants of either 6 m g or 12 mg.

5.2: The NPY mRNA expression of female emus treated with either 110 dexamethasone or saline for 49 days.

5.3: Average daily feed intake per week of female emus treated with 112 either saline or dexamethasone for 49 days.

5.4: Live weight of female emus treated with either saline or 113 dexamethasone for 49 days, on Day -14, Day 14 and Day 49.

5.5: Serum insulin concentration of female emus treated with either 115 saline or dexamethasone for 49 days.

5.6: Blood glucose concentration of female emus treated with either 116 saline or dexamethasone for 49 days.

5.7: Serum thyroxine concentration of female emus treated with 117 either saline or dexamethasone for 49 days.

5.8: Serum triiodothyronine concentration of female emus treated 118 with either saline or dexamethasone for 49 days.

5.9: T3/T4 ratio of female emus treated with either saline or 119 dexamethasone for 49 days.

5.10: Serum luteinising hormone concentration of female emus 120 treated with either saline or dexamethasone for 49 days.

5.11: Serum corticosterone concentration of female emus treated with 121 either saline or dexamethasone for 49 days.

6.1: The NPY and VIP mRNA expression of incubating and 132 non-incubating male emus determined using the R P A technique.

6.2: The serum prolactin concentration from Week 3 to Week 6 of 134 incubation of incubating and non-incubating male emus.

6.3: Mean blood glucose concentrations from Week 3 to Week 6 of 135 incubation of incubating and non-incubating male emus.

6.4: Mean serum glucagon concentrations from Week 3 to Week 6 of 136 incubation of incubating and non-incubating male emus.

6.5: Mean serum insulin concentrations from Week 3 to Week 6 of 137 incubation of incubating and non-incubating male emus.

IX

Figures (continued)

Page

6.6: Mean serum insulin/glucagon ratio from W e e k 3 to Week 6 of 138 incubation of incubating and non-incubating male emus.

6.7: Mean serum triiodothyronine concentrations from Week 3 to Week 139 6 of incubation of incubating and non-incubating male emus.

6.8: Mean serum thyroxine concentrations from Week 3 to Week 6 140 of incubation of incubating and non-incubating male emus.

6.9: The T3/T4 ratio from Week 3 to Week 6 of incubation of incubating 141 and non-incubating male emus.

6.10: Mean serum corticosterone concentration from Week 3 to Week 6 142 of incubation of incubating and non-incubating male emus.

7.1: The sampling and measurement schedule for the pre-treatment and 149

treatment periods.

A3.1: Correlation analysis of the relationship between fat score and 214 percentage total adiposity of female emus in experiment 2 (saline treated = open circles and dexamethasone treated = solid circles).

A3.2: Liver tissue from female emus following chronic treatment with 216 either saline (left) or dexamethasone (right).

A3.3: Correlation analysis of the relationship between NPY gene expression 220

relative to P-actin and corticosterone concentration (non-incubators = open circles and incubators = solid circles).

Tables

2.1: Tissues weighed for each experiment. 32

2.2: Assay statistics. 51

2.3: The intra-assay coefficients of variation for the hormone assays 51 performed.

2.4: RNA polymerase and restriction enzymes required for each probe 54

to obtain sense and antisense strands.

2.5: Reagents for RPA hybridisation. 56

3.1: Volumes of reagents used in the reverse transcription reactions. 62

3.2: Volumes of reagents used in the polymerase chain reactions. 63

Tables (continued)

Page

3.3: Thermal cycling conditions used for the polymerase chain reaction. 63

3.4: Volumes of reagents used in nested PCR. 64

3.5: Volumes of reagents used in the ligation reaction. 65

3.6: Volumes of reagents used for colony PCR. 66

3.7: Volumes of reagents used for restriction digests of plasmids 68 containing chicken or mammalian leptin fragments.

3.8: The thermal cycling conditions for the Marathon RACE reactions. 72

3.9: Leptin receptor induction of the chicken leptin bioassay by emu 75

and chicken serum samples.

3.10: Leptin receptor induction of the chicken leptin receptor bioassay 76 by emu serum samples collected over the course of year.

4.1: Composition of the low and high fat diets. 82

4.2: Macronutrient and energy content of the low and high fat diets. 82

4.3: Total feed, protein, fibre, fat and energy intakes over 68 days 84 (includes pre-treatment period) of male emus fed low and high fat diets.

4.4: The fat content of muscle and liver tissue from male emus fed 87

either low or high fat diets for 54 days.

4.5: The body composition of male emus receiving either low or high 87

fat diets for 54 days.

5.1: The mRNA expression of NPY relative to G-actin of female emus 110

treated with either dexamethasone or saline for 49 days.

5.2: The carcass composition of female emus treated with either 111

dexamethasone or saline for 49 days.

5.3: The fat scores of female emus on Day 0 and Day 49 of treatment 114

with either dexamethasone or saline.

6.1: The mRNA expression of NPY and VIP relative to p-actin of 132

incubating and non-incubating male emus.

6.2: The total feed intake over Week 6 of incubation and the body 133 composition of incubating and non-incubating male emus at the end of

Week 6 of incubation.

XI

Tables (continued)

Page

7.1: The m R N A expression of N P Y and VIP in starved and control 152

male emus.

7.2: The average daily feed intake of male and female emus in control 152 and starved groups one week before, and during a subsequent one-week

treatment period.

7.3: The live weight of male and female emus in the control and starved 153 group before and after a 7 day treatment period.

7.4: The body composition of control and one week starved male emus. 153

7.5: Serum insulin concentrations of male and female emus in each 154 group before and after a 7 day treatment period.

7.6: Serum glucagon concentrations of male and female emus in each 154 group before and after a 7 day treatment period.

7.7: The insulin/glucagon ratio of male and female emus in each group 155 before and after a 7 day treatment period.

7.8: Serum thyroxine concentrations of male and female emus in each 155 group before and after a 7 day treatment period.

7.9: Serum triiodothyronine concentrations of male and female emus in 156 each group before and after a 7 day treatment period.

7.10: Serum T3/T4 ratio of male and female emus in each group before 156 and after a 7 day treatment period.

7.11: Serum corticosterone concentrations of male and female emus in 157 each group in each group before and after a 7 day treatment period.

7.12: Serum P-hydroxybutyrate concentrations of male and female emus 157 in each group in each group before and after a 7 day treatment period.

7.13: Serum urea concentrations of male and female emus in each group 158 in each group before and after a 7 day treatment period.

A3.1: The mRNA expression of NPY and VIP relative to p-actin of 219 incubating (n=3) and non-incubating (n=6) male emus.

A3.2: Correlation analysis comparing the expression of NPY relative to 220

P-actin in incubating and non-incubating male emus with corticosterone

and prolactin concentrations and the insulimglucagon ratio.

1

General Introduction

General Introduction 2

The emu has evolved in a harsh environment and is able to contend with extremes of

climate and food availability. A unique feature of the emu is its ability to fatten during the

summer, a time when individual birds can dramatically increase their food intake and fat

depot. As the oil produced from emu fat has proven therapeutic qualities there is the

potential for emu farming to become a successful commercial venture. However, this

potential is limited by the seasonal variation in appetite and subsequently adiposity, which

prevents consistent market supplies of emu products. The industry could overcome these

limitations if methods were developed to limit the decrease in food intake and reduction in

the fat depot over the breeding season.

In other species, mainly mammalian, an integrated model for the control of energy balance

has been developed. Metabolic hormones including, insulin, glucagon, thyroxine and

triiodothyronine, and leptin, from adipose tissue, are peripheral factors that control energy

balance through their effects on food intake and metabolic rate. One way that metabolic

hormones keep energy intake and energy expenditure equal is by acting on neuropeptides

involved in regulating appetite and satiety within the hypothalamus. One of the major

neuropeptides involved in appetite control is neuropeptide Y (NPY), and one of the major

regulators of N P Y gene expression is leptin. The control of appetite and adiposity by

N P Y , leptin and peripheral hormones has not been investigated in the emu. As such, w e

do not know how well this model for the control of energy balance applies to the emu.

The aim of this thesis has been to improve the understanding of the control of appetite and

adiposity in the emu. To address this aim, the general hypothesis of this thesis was that

seasonal variation in appetite and adiposity in the emu operates under the general model

developed in other species. The success of this study depends on how well the emu

adheres to the general model for the control of appetite and adiposity. Given the strong

evolutionary pressure for efficient use of available energy sources, the emu might have

adopted particular strategies to control its energy balance. However, by comparing the

emu to other species these strategies will be identified. In addition, it should be possible to

develop methods for successfully manipulating appetite and adiposity to improve

economic returns for the emu producer.

Chapter 1

Literature Review

Page

1. Introduction

2. The emu

3. E m u farming

3.1 History

3.2 Production constraints

3.3 Industry potential

4. Appetite and the regulation of adiposity

4.1 The lipostatic theory

4.2 Control by the hypothalamus

4.3 Fat deposition and mobilization

5. Understanding the control of appetite and adiposity in the emu

5.1 Leptin

5.2 Neuropeptide Y

5.3 Vasoactive intestinal peptide and prolactin

5.4 Metabolic and hormonal consequences of incubation

6. Manipulation of appetite and adiposity

6.1 High fat diets

6.2 Glucocorticoids

6.3 Starvation

7. Conclusions

4

4

6

6

6

8

9

9

9

10

14

14

16

18

19

20

20

22

25

28

Chapter 1 - Literature Review 4

1. Introduction

The biggest problem facing the emu industry is decreasing adiposity due to reduced

appetite during the breeding season. Adipose tissue is the major product of the emu. As

such, improving our understanding of how appetite and fat deposition are controlled will

enable the development of strategies to manipulate and maintain appetite and adiposity for

longer periods of time. In this review I will cover the biology of the emu, and the history,

constraints and potential of emu farming. Following on from this I will review the control

of appetite and adiposity and their manipulation.

2. The emu

Emus belong to a group of flightless birds collectively called ratites. The mature emu

stands up to two metres in height and weighs in the range of 30 to 50 kg. The feathers are

grey-brown in colour, with a lighter ruff around the neck. The face and part of the throat is

generally bare of feathers and grey-blue in colour, with females normally exhibiting darker

colouring (Mawson, 1992). Following pairing, the breeding season commences in M a y

and usually lasts until June, but can continue into October depending on seasonal

conditions such as rainfall.

The onset of the breeding season occurs in response to decreasing day length (Blache &

Martin, 1999; Blache et al, 2001a). The breeding season is associated with a 30-50%

decrease in food intake and weight loss (Blake, 1996) (Fig. 1.1). The weight loss observed

is due to appetite suppression resulting from several causes related to, the onset of the

breeding season, broody behaviour by males while incubating eggs, and the egg laying

period for the females. The female on average lays fifteen eggs in her first season and 22

eggs in subsequent seasons (O'Malley, 1991). The eggs are incubated by the male, or in

incubators for 54 to 56 days (O'Malley, 1991). At the end of the breeding season appetite

increases and weight returns to pre-breeding levels. In the incubating male this is

particularly striking with a reversal from anorexia to a hyperphagic state. The hyperphagic

state enables the males to regain most of the weight lost (24%) within a three-week period

after the cessation of incubation (Van Cleeff, 2001).


Breeding season

1.5

Feed intake (kg/d) 10

0.5-

14

Daylength 12

(h) ~ 10

8

r60

45

30

Dec

Summer

Feb April June

Autumn Winter

Aug

Spring

Dec

Summer

Body Weight (kg)

•—• Female

o—o Male

Figure 1.1: Annual changes in feed intake and live weight of male (open circles) and female (closed'circles) emus (adapted from Blache et al, 2001b).

Chapter 1 - Literature Review °

3. E m u farming

3.1 - History

Commercial emu farming commenced at Kalannie in Western Australia in 1970 (O'Malley,

1993). Although the first farm closed in 1973 it proved to other producers, and the

government, that emus could be used successfully to produce high quality leather. O n the

basis of this success an experimental emu farm was set up at Wiluna by Applied Ecology,

to be run by the Ngangganawili Aboriginal Community as both a tourism and leather

production enterprise (O'Malley, 1993). Following this a private group, Dromaius Pty Ltd

set up an experimental farm at Mount Gibson. The State Government recognised farming

emus as 'technically feasible' in 1987 and approved the sale of 500 emu chicks from the

Wiluna farm (O'Malley, 1993). By 1994 Western Australia boasted 42 operating emu

farms producing emu meat, oil and leather (Smetana, 1994). The growth of the industry

during the 1990's was indicative of the faith placed in it by producers and of the potential

to utilise Australia's native fauna in ecologically sustainable production systems. The

market has experienced a downturn in recent years with the commercial Australian flock

numbering around 50,000 in 1999 (McKinna, 1999). Similar trends have been experienced

in the other countries where producers enthusiastically entered into emu fanning, among

them N e w Zealand, America, China and parts of Europe. The downturn in emu numbers

is the result of poor marketing of emu products (meat, leather and oil) in the wake of

exponential increases in bird numbers, and high production costs.

3.2 - Production constraints

The relative youth of the industry means that much of the information required for a

producer to decide on the best management practices is unknown for the emu. These

unknowns include scientific knowledge of the bird's nutritional requirements, the

heritability of economically significant traits and the correlations between these traits, and

how to manage birds to maximise production. At present the nutritional requirements and

heritability of traits have been based on extrapolation of those for other avian species,

usually the ostrich or chicken (O'Malley, 1996).

Another feature that has contributed to the emu industries problems is the high

reproductive rate of emus, which resulted in an exponential increase in their numbers once

farming commenced. The dramatic increase in bird numbers was problematic because the

development of markets for emu products did not keep pace with the increase in bird

numbers. Marketing groups still remain under a great deal of pressure to develop eager,

Chapter 1 - Literature Review '

receptive markets. To achieve and maintain these markets it is recognised as essential for

producers to reduce the costs of production by maximising productivity (Deeming &

Angel, 1996).

Therefore, maintaining the viability of the emu industry is dependent upon both

establishing stable markets for e m u products and maximising productivity. Maximising

productivity will reduce production costs and allow emu products to be more competitive

in the market place. To maximise the returns per head it is essential to determine the

optimal production conditions and methods for overcoming the utilisation of fat depots due

to breeding behaviour. Furthermore, it will be important to prove to the market that

consistent supplies of all products can be provided.

At present constant supplies of fat throughout the year are prevented by the weight loss that

is associated with the commencement of the breeding season, this in turn limits the

availability of meat and leather (Blake, 1996). In breeding animals the loss of fat is normal

and acceptable. For animals destined for slaughter the loss of fat during the breeding

season is undesirable, particularly when fat prices are high and fat is a significant

component of the economic return obtained from a carcass.

It has been estimated that a 50 week old emu will yield 4kg of fat (Smetana, 1992). Long-

term ex-farm prices have been estimated for the major emu products. Taking into account

the costs of production, a long-term profit of $38 per bird has been estimated (Smetana,

1992). At present these profits are not being realised despite producers receiving, on

average, higher prices for most products than the long-term predictions. The average

prices received for 1999 in Australia were $26.38/L for oil, $62.50 per skin, $11.00/kg for

meat, $4.75 per egg and $28.00 per live bird (Michael, 2000). The potential profits from

emu farming have been eroded by the high cost associated with transport to abattoirs ($10-

$40 per bird) and slaughter ($90-$ 100 per bird), leading to some producers opting to

slaughter on farm and only sell the fat and leather (Michael, 2000). In addition to this

producers have to transport birds on average more than 200kms to the nearest abattoir, as

such they run a serious risk of damage to the birds' skin during transport and a further

reduction in profits (Michael, 2000).

Chapter 1 - Literature Review °

3.3 - Industry potential

The long-term market trends are considered promising based on the public's initial

response and the international interest that has been shown in emu products. The similarity

of the leather produced to that of ostriches and rheas, and the ability to produce it in

commercial quantities, makes targeting the markets for these other leathers a logical action.

Other products such as carved eggs and jewellery crafted from feathers and nails are aimed

at the tourist market, making their future returns difficult to estimate.

Emu meat makes up 30% of the carcass and is sold domestically and internationally

(Dingle, 1997). It is a game meat derived from the thigh and pelvic area and has many

healthy attributes (Berge et al, 1997). These include low levels of fat, cholesterol and

calories (Dingle, 1997). In recent years emu meat has been awarded the Australian Heart

Foundation tick of approval, a valuable and marketable endorsement. In addition, the fat

that is present in emus is predominantly oleic acid, a mono unsaturated fatty acid, which is

beneficial to human health (Dingle, 1997).

About 50% of the profits received from an emu come from fat, making it an important

component of the carcass, especially as it makes up only 17.5% of the live weight (Dingle,

1997). The major consumers of fat are cosmetics companies, which use it as a base for

cosmetics and therapeutic agents. Furthermore, the reputed therapeutic properties of emu

oil have seen it accepted as an alternative, natural treatment for muscle and joint pain. The

demand for emu oil for this purpose is likely to continue, particularly now that scientific

support for its therapeutic properties has been obtained (Snowden & Whitehouse, 1997;

Whitehouse et al, 1998; Lopez et al, 1999). The scientific support for emu oils therapeutic

properties has also allowed an application to be made for therapeutic goods approval (now

pending) that will help to ensure the future of this product and the emu industry.

The problem of seasonal cycles of adiposity still remains a constraint to producers because

it narrows the window of opportunity for slaughtering the birds. If this can be overcome

emu farmers will have a better chance to establish markets for all their products and

improve their profit margins. To do this it is essential that w e increase our understanding

of the control of appetite and fat deposition in the emu so that strategies to manipulate

appetite and adiposity can be developed. To date no information exists regarding the

control of appetite and adiposity in the emu. However, the literature contains a wealth of

Chapter 1 - Literature Review

information on the control of appetite and adiposity in other species and this can be used to

identify a starting point for research in the emu.

4. Appetite and the regulation of adiposity

4.1 - The lipostatic theory

It has been proposed that an indirect calorimetry mechanism exists that regulates adipose

reserves via sensitivity of the hypothalamus to the concentration of circulating metabolites

(Kennedy, 1953). The lipostatic theory was developed on the basis of observations with

rats that demonstrated they adjusted their food intake to very accurately maintain a constant

fat mass and calorie intake (Adolph, 1947; Kennedy, 1950).

This theory was further supported by force-feeding experiments. Intragastrically

cannulated dogs, fed diets corresponding to below, at and above their own determined

requirements, adjusted their food intake such that the calorie intake was constant, and

where possible at the original level (Janowitz & Hollander, 1955). Force-feeding rats and

guinea pigs for prolonged periods above their requirements resulted in the development of

obesity, whereas starved animals lost weight (Harris et al, 1986; Conn & Joseph, 1962;

Harris & Martin, 1984a & 1984b). In addition, when animals returned to ad libitum

feeding, their food intake and body weight returned to the original level. This indicates

that the regulation of food intake and appetite occurs via recognition of the fat store size

and modification of food intake to maintain it at a "set point".

4.2 - Control by the hypothalamus

The lipostatic theory, assumes that there are sites in the brain that receive signals pertaining

to energy reserves and co-ordinate a response in terms of appetite, energy expenditure and

metabolism. The hypothalamus is thought to be this site. Research has shown that damage

to the hypothalamus of mammalian and avian species results in hyperphagia and

consequently obesity (Brobeck, 1946; Lundbaek & Stevenson, 1947; Hervey, 1959; Anand,

1961; Lepovsky & Yasuda, 1965; Smith, 1968). In addition, decreased energy expenditure

often contributes to the development of obesity in these animals as evidenced by a decline

in the level of activity (Hetherington & Ranson, 1942; Brobeck, 1946).

The ventromedial region of the hypothalamus has been proposed as the site of action for

appetite and adiposity control in the hypothalamus. This is based on the observations of

Hetherington & Ranson (1940) that of discrete, localised lesions placed in almost all parts

Chapter 1 - Literature Review 1 U

of the hypothalamus, only those in the tuberal or posterior regions, and most effectively in

the region of the ventromedial nuclei, induced obesity. Other research where obesity has

been induced by creating bilateral lesions in the ventromedial region of the hypothalamus

have supported Hetherington and Ranson's findings (1940), showing that such lesions

cause the highest levels of obesity, primarily via increased food intake (Hetherington &

Ranson, 1942; Brobeck, 1946; Kennedy, 1950; Hervey, 1959).

The findings of Brobeck (1946), Kennedy (1950) and Hervey (1959) indicated that

initially, animals with lesions had an insatiable appetite that diminished with time but still

remained above that of their intake prior to the operation. In the lesioning experiments of

Hetherington & Ranson (1942) obesity occurred without increases in food intake. Instead,

some animals returned to their previous level of food intake but displayed lower levels of

activity than recorded prior to the procedure. This would indicate that the hypothalamus,

as well as controlling appetite, might have some role in determining the energy expenditure

of an animal based on its energy stores. There is also evidence that damage to the

hypothalamus induces obesity via a shift in metabolism to favour fat deposition. For

example, Van Putten et al (1955) pair fed rats with hypothalamic lesions to the level of

unlesioned controls, which resulted in the pairs having identical body weights. However,

carcass composition analysis revealed significantly higher proportions of fat in the lesioned

animals.

4.3 - Fat deposition and mobilisation

The preceding sections have demonstrated that the size of the fat depot is regulated by the

hypothalamus to maintain energy reserves at an appropriate level or "set point". W h e n the

energy reserve is too low, appetite is increased and adipose sparing mechanisms are

initiated, and when the energy reserve exceeds requirements the animal feels sated and

mechanisms to increase energy expenditure may be initiated. Research since the

proposition of the lipostatic theory has demonstrated that these mechanisms rely on the

interaction of hormones, metabolites and signals from the digestive tract (Fig. 1.2).

In the gastrointestinal tract enzymes convert carbohydrates to monosaccharides, fats to fatty

acids and glycerol and proteins into amino acids (Duke, 1986). These enzymes are released

in response to food consumption and digestion. Cholecystokinin, a hormone originating

from the upper small intestine, is released in response to duodenal distension, vagal

stimulation and the presence of nutrients from food (Duke, 1986). Elevation of


cholecystokinin concentrations decreases food intake via central actions (Savory & Gentle,

1980; Denbow & Myers, 1982; Denbow, 1986).

Nutrients absorbed from the gastrointestinal tract enter the hepatic portal blood system and

are transported to the liver. Elevations of glucose, amino acid or fatty acid concentrations

decrease food intake (Denbow, 1999). Glucose decreases food intake via chemoreceptors

in the liver that act on the central nervous system (Forbes, 1988). The lipid component of

the absorbed nutrients is converted to very low-density lipoproteins (VLDL's) in the liver,

and transported to peripheral tissues (Stevens, 1996). Lipoprotein lipase hydrolyses the

VLDL, releasing fatty acids that are taken up by tissues to meet their energy requirements

or stored in adipose tissue (Stevens, 1996).

\_ Parasympathetic (Vagus) J

CCK

Food Intake

Digestive tract

nutrients

Adrenal gland

Liver

~r lipogenesis

Catecholamines

Glucocorticoids

Insulin

T Pancreas

gluconeo genesis

tissue

Glucose

• - lipolysis -| FFA

ucagon

Leptin

Figure 1.2: A model for the control of food intake and adipose tissue deposition. T h e blue lines indicate

stimulation (solid) or inhibition (dashed) of hormone release or production. T h e black and red lines indicate

negative (dashed) or positive (solid) effects. C C K = cholecystokinin, C R H = corticotrophin-releasing

hormone, F F A = free fatty acid, N P Y = neuropeptide Y .

The storage or mobilisation of energy reserves is controlled by the interactions of

hormones. The most important hormones in the storage and mobilisation of energy

reserves are insulin, glucagon, glucocorticoids, catecholamines and leptin. In mammals,

11

Chapter 1 - Literature Review t ̂

insulin is the primary regulator of metabolism. Insulin stimulates the uptake and storage of

lipids. In avian species, it does this by acting on the liver to increase glucose uptake, inhibit

gluconeogenesis, and increase lipogenesis and the release of V L D L from hepatocytes

(Stevens, 1996). In avian adipose tissue, insulin increases glucose uptake, inhibits free

fatty acid release, stimulates the conversion of glucose to fat and stimulates lipoprotein

lipase concentrations (Griminger, 1986; Hazelwood, 1986). These actions of insulin spare

the triglyceride stores and increase adipose deposition. In mammals, insulin acts centrally

to decrease appetite via suppression of neuropeptide Y (NPY) gene expression (Schwartz

et al, 1991 & 1992). It is likely that this also occurs in avian species, and provides a

negative feedback loop to maintain energy reserves at the correct level.

In contrast to mammals, glucagon is the primary regulator of metabolism in avian species.

Glucagon mobilises energy reserves by inhibiting lipogenesis and stimulating

gluconeogenesis in the liver and stimulating lipolysis in adipose tissue (Stevens, 1996).

The increased lipolytic activity of adipose tissue releases free fatty acids into the peripheral

circulation (Braganza et al, 1973). Glucagon release from the pancreas is stimulated by

insulin, free fatty acids and cholecystokinin and profoundly inhibited by glucose, providing

positive and negative feedback loops (Hazelwood, 1986).

Catecholamines are released from the adrenal gland in response to signals from the

sympathetic nervous system (Stevens, 1996). The catecholamines mobilise stored energy

reserves primarily from the liver, and to a lesser extent from adipose tissue, by stimulating

lipolysis and inhibiting fatty acid synthesis (Stevens, 1996). Glucocorticoids are released

from the adrenal gland in response to increasing adrenocorticotropin ( A C T H )

concentrations in the anterior pituitary (Stevens, 1996). The increase in the concentration

of A C T H is in turn controlled by corticotrophin-releasing hormone (CRH) at the

hypothalamic level (Laycock & Wise, 1996; Kacsoh, 2000). The glucocorticoids have

opposing actions in liver and adipose tissue, stimulating lipogenesis and gluconeogenesis

in the liver, and lipolysis in adipose tissue (Griminger, 1986; Stevens, 1996).

Glucocorticoids also have a negative feedback function, suppressing hypothalamic C R H

production (Kacsoh, 2000). Elevations in C R H , as well as increasing glucocorticoid

release, also suppress the actions of N P Y to increase appetite (McCarthy et al, 1993;

Menzaghi et al, 1993). This action of C R H on N P Y is proposed as one of ways that

depressed appetite is maintained in humans suffering from anorexia nervosa (St0ving et al,

1999).


C R H is regulated by leptin, a hormone secreted by adipocytes (section 5.1). Leptin

increases the hypothalamic C R H content and expression in vivo, and C R H secretion in

vitro (Schwartz et al, 1996a; Costa et al, 1997; Uehara et al, 1998). In rats, pre-treatment

with antibodies against C R H attenuates the ability of leptin to decrease appetite (Okamoto

et al, 2001). As such, C R H may be involved in the suppression of N P Y expression and

appetite following central leptin administration (Jang et al, 2000). The role of leptin in

avian metabolism has not been extensively studied and is complicated by doubts over the

veracity of published sequences for the chicken leptin gene (see section 5.1 for further

discussion). The evidence to date indicates that leptin expression in the liver, but not

adipose tissue, increases in response to insulin administration (Ashwell et al, 1999a;

Richards et al, 1999). In mammals, the expression of the leptin gene, and leptin secretion,

are stimulated by insulin (Saladin et al, 1995; Barr et al, 1997). Insulin and leptin both act

centrally to decrease appetite (Mistry et al, 1997; Tang-Christensen et al, 1999).

There is evidence that leptin increases the sensitivity of animals to the satiety effects of

cholecystokinin (Emond et al, 1999; Matson & Ritter, 1999). Furthermore, leptin increases

the metabolic rate and energy expenditure (Pelleymounter et al, 1995; Collins et al, 1996;

H w a et al, 1996; H w a et al, 1997; Mistry et al, 1997). Both metabolic rate and energy

expenditure are disrupted, along with appetite, following hypothalamic lesions

(Hetherington & Ranson, 1942; Brobeck, 1946; Van Putten et al, 1955). As such, leptin is

a powerful regulator of adipose reserves at the hypothalamic level, via its ability to alter

appetite, energy expenditure and metabolism, in response to changes in energy balance.

The action of leptin to alter appetite, energy expenditure and metabolism at the

hypothalamic level is mediated by the interaction of a large number of neuropeptides.

N P Y is one of these neuropeptides, it has been mentioned above and has been investigated

in the course of this thesis. N P Y is discussed further in section 5.2. Other hypothalamic

neuropeptides that have a role in the control of appetite, energy expenditure and

metabolism include agouti-related protein, cocaine and amphetamine-regulated transcript,

orexin/hypocretin, proopiomelanocortin and C R H (McCarthy et al, 1993; Menzaghi et al,

1993; Mizuno et al, 1997; Ollmann et al, 1997; Sakurai et al, 1998; Heinrichs & Richard,

1999; Vrang et al, 1999). However, these neuropeptides were not investigated in the

course of this thesis. Instead, m y investigations have focused on N P Y as it has been

Chapter 1 - Literature Review lH

extensively studied in avian species and is considered to be a major central pathway

involved in the control of appetite, energy expenditure and metabolism.

5. Understanding the control of appetite and adiposity in the emu

Hormones, metabolites and genes interact to control appetite via their response to food

intake, absorption of nutrients and changes in the size of the adipose depot (Fig. 1.2). In

the following sections those hormones, metabolites and genes that were investigated in the

course of this thesis are discussed further.

5.1 - Leptin

In the last decade the leptin gene has been cloned and sequenced for a number of species

(Zhang et al, 1994; Masuzaki et al, 1995; Murakami & Shima, 1995; Ramsay et al, 1998).

The gene encodes a 4.5 kilobase adipose tissue specific messenger R N A ( m R N A ) in both

mice and humans (Zhang et al, 1994; Masuzaki et al, 1995). There is 8 4 % homology

between the human and mouse leptin genes. The product of this gene, leptin, is a satiety

signal involved in the regulation of appetite and adiposity (Friedman, 1996; Sinha, 1997;

Auwerx & Staels, 1998; Buchanan et al, 1998; Inui, 1999b). The concentration of leptin

corresponds closely to body mass index (BMI), a correlate of adiposity (Maffei et al, 1995;

Considine et al, 1996; McGregor et al, 1996; Rosenbaum et al, 1996; Schwartz et al,

1996b; Niskanen et al, 1997; Tataranni et al, 1997; Parry et al, 1998; Jensen et al, 1999).

In non-human species leptin concentrations and leptin m R N A expression are directly

correlated with adiposity (Frederich et al, 1995; Maffei et al, 1995; D e Schepper et al,

1998; Hissa et al, 1998; Robert et al, 1998).

Zhang et al (1994) hybridised the leptin gene to Southern blots of genomic DNA from the

mouse, rat, rabbit, vole, cat, cow, sheep, pig, human, chicken, eel and fruit fly. For all the

vertebrate species, detectable signals were present, indicating that they contained D N A

homologous to the mouse leptin gene. The presence of a signal in the chicken indicates

that the leptin gene is likely to be present in other avian species, including emus.

The controversy surrounding the cloning of the avian leptin gene.

Support for the presence of the leptin gene in avian species has been obtained from the

sequencing of the chicken leptin receptor (Horev et al, 2000). T w o groups also claim to

have cloned the chicken leptin gene (Taouis et al, 1998; Ashwell et al, 1999a & 1999b).

Subsequently, a recombinant chicken leptin has been produced (Raver et al, 1998). It is

Chapter 1 - Literature Review I-5

claimed that the chicken sequence shares a high degree of homology (95-97%) with the

mouse leptin gene (Taouis et al, 1998; Ashwell et al, 1999a). However, the chicken leptin

receptor shares only a low degree of homology with mammalian sequences (on average

60%), and this casts doubt on the claims to have sequenced the chicken leptin gene as

receptor sequences are normally more highly conserved than the gene (Horev et al, 2000).

Further doubt is cast by the maintenance of appetite in chickens receiving central injections

of the highly homologous (95-97%) murine recombinant leptin (Bungo et al, 1999). This

is an unexpected finding given that human recombinant leptin, despite being less

homologous to the murine sequence (84%), can decrease appetite when administered to the

mouse (Hwa et al, 1996). Southern hybridisation has demonstrated that the homology

between mouse and chicken leptin genes cannot be more than 72-77% (Varma, 2000).

This degree of homology is much lower than the 95-97% homology at the nucleotide level

claimed by Ashwell et al (1999a) and Taouis et al (1998). Northern hybridisation with a

mouse leptin probe to chicken adipose R N A failed to produce a signal, and P C R and RT-

P C R using primers designed for the mouse and the published chicken leptin sequences also

failed to generate P C R products (Friedman-Einat et al, 1999). Subsequently, claims to

have successfully cloned the chicken leptin gene have been openly challenged by avian

researchers (Dunn et al, 2001; Varma, 2000; Horev et al, 2000; Friedman-Einat et al,

1999).

Mechanism of action of leptin

Daily administration of recombinant leptin to mice lacking functional leptin (ob/ob

genotype) causes time and dose dependent reductions in body weight and food intake

(Weigle et al, 1995; Tartaglia et al, 1995; Pelleymounter et al, 1995; Halaas et al, 1995;

Campfield et al, 1995; Stephens et al, 1995; H w a et al, 1996). In normal mice,

recombinant leptin reduces food intake, even after 24 hours of fasting (Pelleymounter et al,

1995; Rentsch et al, 1995; Mistry et al, 1997). The action of leptin to decrease body

weight does not occur purely via appetite suppression, as the food intake of normal mice

treated with leptin returned to its original level without their body weight returning to pre-

injection levels. Other changes attributed to leptin administration include alterations in

metabolism, level of activity, oxygen consumption, and the normalisation of body

temperature, serum insulin and glucose levels in the ob/ob mouse (Pelleymounter et al,

1995; H w a et al, 1996; H w a et al, 1997; Mistry et al, 1997).

Collins et al (1996) proposed that leptin regulates the extent of adipose tissue reserves by a


mechanism that involves both appetite suppression and sympathetic nervous system

signaling to increase thermogenesis and energy expenditure in brown adipose tissue as fat

mass increases. To test this hypothesis leptin was administered to ob/ob mice and their

noradrenaline turnover was measured after blocking catecholamine synthesis, which is

indicative of the sympathetic outflow. The administration of leptin caused a selective

increase in noradrenaline turnover in brown adipose tissue, with no effect upon turnover in

white adipose tissue. Taken together these findings demonstrate that leptin regulates body

weight via effects on both food intake and the level of energy expenditure.

5.2 - Neuropeptide Y

Leptin's role in the regulation of energy reserves is as a signal of the adipose depot size,

that initiates appropriate responses in terms of appetite, energy expenditure and metabolic

rate in response to deviations from the "set point". One way in which leptin achieves this is

via its effects on neuropeptide Y (NPY), a potent appetite stimulant. There is an inverse

relationship between the expression of leptin and N P Y .

Tatemoto et al (1982) isolated NPY, a 36 amino acid peptide, from porcine brain extracts.

In both birds and mammals, N P Y is widely distributed in the central nervous system

(Tatemoto et al, 1982; Allen et al, 1983; Chronwall et al, 1985; Boswell et al, 1998;

Gehlert, 1999; Inui, 1999a; W a n g et al, 2001). The highest levels of expression occur in

the hypothalamus (Adrian et al, 1983; Allen et al, 1983; Chronwall et al, 1985; Kameda et

al, 2001). N P Y is a potent central appetite stimulant in avian and mammalian species

(Levine & Morley, 1984; Stanley & Leibowitz, 1984 & 1985; Stanley et al, 1985a & 1986;

Kuenzel et al, 1987; Paez & Myers, 1991; Richardson et al, 1995; Choi et al, 1997; Woods

et al, 1998; Inui, 1999a). N P Y has a pivotal role in the control of appetite and it has been

proposed that increased appetite occurs via NPY's actions at the level of the hypothalamus

in both avian and mammalian species (Stanley et al, 1993; Stanley et al, 1985a; Strader &

Buntin, 2001).

The food intake of animals is higher once food is returned after a period of fasting. This

mirrors the increase in feed intake following central N P Y administration, and research

indicates that endogenous N P Y mediates the feeding responses provoked by food

deprivation (Stanley et al, 1992; Bivens et al, 1998). In addition, it has been found that

passive immunisation against N P Y markedly suppresses feed intake in non-fasted rats

(Dube et al, 1994). It is apparent from these results that N P Y is an important signal


stimulating ingestive behaviour.

In addition to its effects on appetite, NPY promotes adipose deposition by decreasing

energy expenditure and thermogenesis, and increasing the efficiency of energy storage

(Roscoe & Myers, 1991; Billington et al, 1994; Currie & Coscina, 1995; Stephens et al,

1995; Woods et al, 1998). These actions of N P Y decrease the metabolic rate but do not

appear to involve modulation of the thyroid, as N P Y injection has no effect on thyroid

gland weight, or the serum concentrations of thyroid stimulating hormone, triiodothyronine

and thyroxine (Malendowicz & MiSkowiak, 1990). N P Y does however increase the

consumption of carbohydrate preferentially to fats or protein (Stanley et al, 1985b; Morley

et al, 1987). N P Y also increases the respiratory quotient, indicating that carbohydrate

catabolism is increased (Menendez et al, 1990). As a consequence of this, fat synthesis is

increased.

NPY also interacts with metabolic hormones to promote adipose deposition and energy

sparing mechanisms. Elevations in N P Y expression increase the level of circulating

insulin (Walker & Romsos, 1993; Billington et al, 1994; Tomaszuk et al, 1996; Parikh &

Marks, 1997). This favours fat deposition via increased triglyceride synthesis and

deposition in white adipose tissue (Stevens, 1996). As such, prolonged elevation of N P Y

induces obesity, via increased appetite and metabolic changes that favour fat synthesis and

deposition (Stanley et al, 1986). The effect of N P Y to increase the concentration of insulin

may also function as a negative feedback loop reducing N P Y expression and preventing

excessive adipose deposition. This idea is supported by research demonstrating that the

central administration of insulin decreases N P Y m R N A expression in rats (Schwartz et al,

1991 & 1992). In rats, without access to food, the central administration of N P Y increases

the concentration of glucagon and causes a transient increase in plasma glucose without

effecting the concentration of corticosterone (Marks & Waite, 1996). Central N P Y

administration also decreases insulin sensitivity (Marks & Waite, 1997). During fasting,

the effects of N P Y to increase the concentration of glucagon and decrease insulin

sensitivity interfere with the control of glucose metabolism by insulin. Hypothalamic N P Y

content is rapidly increased in avian and mammalian species by food deprivation or

restriction (Chua et al, 1991; Sahu et al, 1992; Marks et al, 1992; McShane et al, 1993;

Davies & Marks, 1994; Stricker-Krongrad et al, 1997; Jang & Romsos, 1998; Boswell et

al, 1999; Mizuno etal, 1999; Boswell, 2001; Kameda et al, 2001). This elevation in N P Y

is associated with decreased insulin and glucose and increased corticosterone

Chapter 1 - Literature Review 1 0

concentrations (Marks et al, 1992; Mercer et al, 1996; Jang & Romsos, 1998; Dallman et

al, 1999; Mizuno etal, 1999).

Role of NPY in the seasonal cycle of food intake

In some species seasonal cycles of food intake and fat metabolism are observed. These are

proposed to occur via a "sliding set point" for homeostasis (Mrosovsky, 1976; Mrosovsky

& Sherry, 1980; Mercer, 1998). The mechanisms that control this are not well understood,

and the role of N P Y is not clear. In sheep, lower levels of N P Y expression are associated

with periods of low appetite (Clarke et al, 2000). Siberian hamsters, a hibernating species,

also have lower levels of N P Y m R N A expression during short photoperiods, when appetite

is low (Reddy et al, 1999). In addition, this experiment demonstrated that 48 hours of food

deprivation, during both short and long photoperiods, increased N P Y m R N A expression

(Reddy et al, 1999). Furthermore, the Siberian hamster increases its food intake in

response to the central administration of N P Y , on both long and short photoperiods (Boss-

Williams & Bartness, 1996). Another hibernating species, the golden mantled ground

squirrel, also increases its food intake in response to central N P Y infusion (Boswell et al,

1993). It can be inferred from these findings that N P Y expression is correlated with

appetite, and that the responsiveness of central pathways to NPY's actions are not impaired

by changes in photoperiod.

It is likely that the "sliding set point" for homeostasis applies to avian species as well, via

similar mechanisms. In the white-crowned sparrow, as with the Siberian hamster, central

administration of N P Y increases food intake on both short and long days (Richardson et al,

1995). This experiment also demonstrated that during long days, when food intake is

increased, the animals were more sensitive to the effects of N P Y on appetite (Richardson et

al, 1995). As such, a plausible mechanism for photoperiod induced changes in the "sliding

set point" for homeostasis is altered sensitivity to central appetite mediators.

5.3 - VIP, prolactin and reproduction in birds

Changing photoperiod increases the expression of vasoactive intestinal polypeptide (VIP)

and prolactin secretion that are involved in the induction and maintenance of incubation

behaviour in birds. VIP was isolated from porcine small intestine (Said & Mutt, 1970). It

has since been isolated and characterised for a range of species including chickens, cattle,

humans and rats (Nilsson, 1974 & 1975; Carlquist et al, 1979 & 1982; Dimaline et al,

1984). VIP acts as a vasodilator in the periphery, stimulates respiratory chemoreceptors,

Chapter 1 - Literature Review l y

increases cardiac output and induces hypotension and hyperglycemia (Said & Mutt, 1970).

VIP is widely distributed throughout the central nervous system, and is found at highest

levels in the hypothalamus (Bryant et al, 1976; Said & Rosenberg, 1976; Yamada et al,

1982; Kuenzal et al, 1997; Chaiseha & El-Halawani, 1999). Elevations of VIP stimulate

prolactin release in birds and mammals (Kato et al, 197'8; Fuxe et al, 1979; Macnamee et

al, 1986; Opel & Proudman, 1988; Sharp et al, 1989; Pitts et al, 1994; Maney et al, 1999;

Vleck & Patrick, 1999). In avian species the rise in VIP, and subsequently prolactin, is

involved in the induction and maintenance of broody behaviour (El Halawani et al, 1995;

Crisostomo et al, 1997; Bedecarrats et al, 1999; Sockman et al, 2000). In birds immunised

against VIP, photo-stimulation fails to induce a rise in prolactin or induce incubation

behaviour, demonstrating that VIP is an essential intermediary in prolactin response to

photoperiod and for the induction of incubation (El Halawani et al, 1996; Crisostomo et al,

1997).

5.4 - Metabolic and hormonal consequences of incubation

The incubation period in birds can be divided into three phases (Le Maho et al, 1981;

Castellini & Rea, 1992). In the goose, phase I occurs in the first few days of incubation (Le

Maho et al, 1981). Phase I involves a shift from metabolism of ingested nutrients to meet

energy requirements to utilisation of the adipose stores, with minimal weight loss. Phase II

from day 3 to 30 of incubation is associated with a reduced resting metabolic rate and

complete reliance on adipose stores for energy requirements. The third phase from day 30

until the completion of incubation involves protein breakdown to supply the animals

energy requirements as the lipid stores are depleted. The hormonal changes associated

with incubation decrease the metabolic rate and spare the animals energy reserves. These

changes include decreased thyroxine and insulin, decreased or unchanged triiodothyronine

concentrations and increased corticosterone and glucagon concentrations (Chieri et al,

1972; Cherel et al, 1988; Groscolas & Leloup, 1989; Van Cleeff, 2001). The incubation

period is also associated with depressed appetite in many avian species (Mrosovsky, 1976;

Savory, 1979; Mrosovsky & Sherry, 1980; Zadworny et al, 1985; Cherel et al, 1988).

Prolactin is elevated during incubation and has been shown to have variable effects on

appetite. In mammalian species, food intake can be increased, decreased or remain

unchanged when prolactin is suppressed (Shani et al, 1975; Brien, 1986; Bartness et al,

1987; Curlewis etal, 1988 & 1991; Salah et al, 1995). In both mammalian and avian

Chapter 1 - Literature Review ^

species, appetite increases, decreases or does not change in response to prolactin

administration (Shani et al, 1975; Ryg & Jacobsen, 1982; Denbow, 1986; Moore et al,

1986; Bartness et al, 1987; Curlewis et al, 1988; Buntin & Figge, 1988; Gerardo-Gettens et

al, 1989a &1989b; Heil, 1999; Ebenezer & Parrott, 1991; Byatt et al, 1993; Hnasko &

Buntin, 1993; Noel & Woodside, 1993; Salah et al, 1995; Buntin et al, 1999). In ring

doves the increase in appetite in response to prolactin administration occurs via an increase

in the number of cell bodies that are immunoreactive to N P Y (Strader & Buntin, 2001).

However, despite elevated prolactin concentrations, appetite is not increased during

incubation. It is possible that differential regulation of appetite occurs in the central

nervous system and is dependent on reproductive status, factors such as the egg or nest

stimuli, and the permissive action of other hormones (Buntin & Tesch, 1985; Janik &

Buntin, 1985; Horseman & Buntin, 1995). This differential regulation could take the form

of an inhibition of NPY's response to elevated prolactin levels, or inhibition of NPY's

actions to increase appetite during the reproductive period.

6. Manipulation of appetite and adiposity

The decreased appetite and adiposity observed during incubation is an adaptation to enable

a more important process, namely reproduction, to succeed. It occurs via endogenous

disruption of the mechanisms controlling energy balance. The control of appetite and

adiposity can also be manipulated exogenously, to prevent the animal from maintaining

energy reserves at the correct "set point". Manipulating the "set point" can be achieved

either by physical means (i.e. starvation or force-feeding) or by interfering with the

animal's mechanisms of assessment and response to changes in the extent of the energy

reserves (i.e. hypothalamic lesions, high fat feeding, glucocorticoid treatment). The

manipulations employed during the course of this experiment are discussed below.

6.1 - High fat diets

High fat diets increase adiposity in mammals (Edozien & Switzer, 1978; Faust et al, 1978;

Ausman et al, 1981; Oscai et al, 1984; Diersen-Schade et al, 1985; Storlien et al, 1986;

West et al, 1992; Hill et al, 1992; Boozer et al, 1995; Van Heek et al, 1997; Smith et al,

1998; Kim et al, 2000). This is also observed in chickens (Deaton et al, 1981; Vila &

Esteve-Garcia, 1996). High fat diets induce hyperphagia, although this effect is variable

across species and strains and may be transient (Oscai et al, 1984; West et al, 1992; Hill et

al, 1992; French et al, 1995; Smith et al, 1998).

Chapter 1 - Literature Review ^1

The degree of adiposity induced by a high fat diet is proportional to the diet's fat content

(Edozien & Switzer, 1978; Deaton et al, 1981; Boozer et al, 1995; Vila & Esteve-Garcia,

1996). High fat diets can increase adiposity without increasing energy consumption or

body weight (Oscai et al, 1984; Storlien et al, 1986; West et al, 1992; Boozer et al, 1995;

Smith et al, 1998). They are able to do this because more calories are obtained from fat,

and provided the oxidation of carbohydrates and proteins is sufficient to meet energetic

requirements, fats are stored.

High fat diets increase fat consumption and storage via the following mechanisms:

1. Lower satiety of diets high in fat (French et al, 1995; Covasa & Ritter 1998 &

1999; R o m o n et al, 1999; Havel et al, 1999),

2. Greater absorbance of fat from the diet than protein or carbohydrate (Singh et

al, 1972; Reed et al, 1991; French etal, 1995),

3. Slower induction of fat oxidation from the diet than protein or carbohydrate

(Flattera/, 1985; Hill etal, 1992; Prentice, 1998; Friedman, 1998), and

4. Decreased energy expenditure and heat production in response to high fat diets

(Black et al, 1949, Storlien et al, 1986).

Because of these mechanisms, particularly the lower satiating effect, consumption of high

fat diets can lead to passive over consumption of energy and increased adiposity (Bray &

Popkin, 1998).

In mice, high fat diets induce peripheral, but not central, leptin resistance, and the onset of

resistance is faster the higher the diets fat content (Van Heek et al, 1997). It has been

proposed that leptin is a modulator of weight gain that is overridden by the continuous

presence of high levels of fat in the diet (Ahren, 1999). Such a mechanism allows animals

to take full advantage of high fat food sources when they are available and thereby increase

their chances of surviving subsequent famines. The ability of mice to override the control

of appetite and adiposity by leptin suggests that they have a "thrifty" genotype. The

"thrifty" genotype was proposed by Neel (1955) as an explanation of the greater efficiency

of food utilisation associated with inherited metabolic disorders. These genotypes have

evolved because greater efficiency of food utilisation is an advantage in famine situations

and allows for greater storage of ingested nutrients. However progress has eliminated feast

and famine situations in the Western world and along with the high fat content of today's

diet has rendered this "thrifty" genotype detrimental to health. It seems logical that other

Chapter 1 - Literature Review z z

species have developed similar strategies for coping with feast and famine situations. The

emu is subjected to extremes of food availability in the wild and it is probable that it has

evolved mechanisms to take full advantage of abundant food sources and increase adipose

deposition.

It is likely that these mechanisms may involve metabolic adaptations, however these can

vary widely between species and strains. Rats fed high fat/protein diets had increased

fasting plasma glucose concentrations, glucose intolerance and insulin resistance (Ramirez

et al, 1990). In other experiments, rats fed diets that contained from 12 to 4 8 % of energy

as fat, did not show differences in their insulin or glucose concentrations (Boozer et al,

1995). In mice, glucose and insulin concentrations are unchanged by high fat diets

although insulin sensitivity is reduced (Lemonnier et al, 1975). However, other researchers

have found that high fat diets (58%) increase insulin concentrations, and create a state of

insulin resistance and glucose intolerance (Ahren & Scheurink, 1998).

6.2 - Glucocorticoids

As with high fat diets, the actions of glucocorticoids to increase adiposity result from

hormonal changes that alter metabolism (Fig. 1.3). Glucocorticoid treatment increases

circulating insulin concentrations in avian and mammalian species (Campbell et al, 1966;

Simon, 1984; McKibbin et al, 1992; Corah et al, 1995; Dallman et al, 1995; Berneis et al,

1996; Miell et al, 1996; Makino et al, 1998; Zakrzewska et al, 1999). In birds, increased

concentrations Of insulin favour adipose deposition (Griminger, 1986). A feature of

chronic glucocorticoid treatment is increased adiposity in both avian and mammalian

species (Hausberger & Ramsay, 1953; Dulin, 1956; Nagra & Meyer, 1963; Nagra, 1965;

Simon, 1984; Kafri et al, 1988; Corah et al, 1995). This arises from the induction of insulin

resistance and the action of concomitant elevations of insulin and glucocorticoids to

increase adipose deposition (McKibbin et al, 1992; Dallman et al, 1993; Taouis et al,

1993; Dallman et al, 1995). The increased adiposity observed with glucocorticoid

treatment is not necessarily associated with increased appetite or live weight.

In both birds and mammals, glucocorticoid treatment may be accompanied by decreases in

both appetite and weight gain (Greenman & Zarrow, 1961; Simon, 1984; Kafri et al, 1988;

McKibbin et al, 1992; D e Vos et al, 1995; Makino et al, 1998; Zakrzewska et al, 1999).

As central insulin administration decreases neuropeptide Y (NPY) m R N A expression, the

increase in insulin concentration associated with chronic dexamethasone treatment may

Chapter 1 - Literature Review Z J

contribute to appetite depression (Schwartz et al, 1991; Schwartz et al, 1992). Indeed, in

rats, N P Y was increased in the lateral hypothalamic area following acute but not chronic

dexamethasone treatment (McKibbin et al, 1992). In this experiment insulin was not

altered 4 hours after an injection of dexamethasone, but was increased after 7 days of

treatment and associated with decreased appetite (McKibbin et al, 1992). In dogs, chronic

dexamethasone treatment impairs insulin transport into the central nervous system (Baura

et al, 1996). As such, it would seem unlikely that insulin could suppress hypothalamic

N P Y expression in these animals. In Syrian hamsters, chronic dexamethasone treatment

increased N P Y expression and depressed weight gain without changing the insulin

concentration (Mercer et al, 1996). These conflicting results represent species-specific

differences in response to dexamethasone, and highlight the need for further research in

this area.

The trend for decreased weight gain indicates that glucocorticoids have catabolic effects.

These catabolic effects are particularly pronounced at high doses and result from the

bitonic response of live weight and feed efficiency to glucocorticoids (Devenport et al,

1989; Dallman et al, 1993). This response is due to the presence of two corticosterone

receptors, type I and type II. With increasing glucocorticoid concentrations the type II

receptor is stimulated to a greater extent, promoting tissue catabolism (Devenport et al,

1989). The synthetic glucocorticoid dexamethasone, in contrast to other glucocorticoids,

only stimulates the type II receptor. Despite this dexamethasone treatment can increase

appetite and adiposity (Adams & Sanders, 1992; Corah et al, 1995). This is because the

increase in insulin observed with dexamethasone treatment promotes adipose deposition,

and contributes to increases in the fat to lean mass (Devenport et al, 1989; Corah et al,

1995; Miell etal, 1996; Kolaczynski etal, 1997).

Glucocorticoids have variable effects on glucose concentrations in both avian and

mammalian species (Greenman & Zarrow, 1961; Campbell etal, 1966; Simon, 1984;

Joseph & Ramachandran, 1992; McKibbin et al, 1992; Papaspyrou-Rao et al, 1997;

Makino et al, 1998). In birds, increased concentrations of glucose have a sparing effect on

adipose stores, via decreases in the concentration of glucagon and adipose utilisation

(Hazelwood, 1986). Another feature of glucocorticoid treatment, in both birds and

mammals, is increased liver weight and liver fat contents (Dulin, 1956; Campbell et al,

1966; Adams, 1968; Simon, 1984, Kafri et al, 1988). The increased liver weight is due to

an accumulation of fat in the liver because of the action of glucocorticoids to promote


gluconeogenesis and lipogenesis in this tissue (Dallman et al, 1993). These actions

increase the extent of the energy reserves.

Figure 1.3: The hypothalamic-pituitary-adrenal axis: effects of endogenous (upper diagram) and exogenous (lower diagram) glucocorticoids (solid lines indicate stimulation/increase, dashed lines indicate inhibition/decrease). Grey lines and labels indicate that effects are diminished or unknown. C R H = corticotrophin-releasing hormone, P O M C = proopiomelanocortin (precursor for A C T H ) , A C T H =

adrenocorticotropic hormone. Adapted from Kacsoh (2000).


In the chicken, acute dexamethasone has variable effects on plasma triiodothyronine and

decreases plasma thyroxine concentrations (Mitchell et al, 1986; Darras et al, 1997).

These effects on thyroid hormone concentration are due to inhibition of thyroid stimulating

hormone release, decreases in the release of thyroid releasing hormone (TRH) and a

diminished expression of proTRH m R N A (van Haasteren et al, 1996; Darras et al, 1997).

These changes in thyroid hormones could be expected to decrease the metabolic rate and

have a sparing effect on adipose stores. Indeed acute dexamethasone treatment decreases

heat production in chickens, indicating a decreased metabolic rate (Mitchell et al, 1986).

Glucocorticoids also impact on reproductive function in birds. In chickens, corticosterone

administration induced ovarian regression and decreased the concentration of luteinising

hormone, progesterone and oestradiol (Etches et al, 1984). Acute dexamethasone

treatment decreased luteinising hormone secretion and prevented ovulation when given 13-

14 hours before the expected time of ovulation (Soliman & Huston, 1974; Wilson &

Lacassagne, 1978). This effect is due to the action of dexamethasone to inhibit A C T H

release and suppress corticosterone secretion (Arimura et al, 1969; Sirett & Gibbs, 1969;

Buckland et al, 191 A; Etches, 1976; Arvat et al, 1998). The suppression of corticosterone

secretion prevents the positive feedback of corticosterone on luteinising hormone secretion

(Wilson & Lacassagne, 1978).

Clearly, manipulations of the hypothalamic-pituitary-adrenal axis (HPA) by glucocorticoid

administration can alter the metabolic rate and hormone concentrations to favour increases

in adiposity. As such, treating emus with glucocorticoids provides an opportunity to

improve our understanding of the control of appetite and metabolism, while determining

the potential of glucocorticoids as a means of increasing adiposity in the emu.

6.3 - Starvation

In contrast to treatment with high fat diets or glucocorticoids, the effects of starvation on

appetite and adiposity are clear. Starvation increases appetite, decreases adiposity and

initiates mechanisms to conserve energy reserves. These mechanisms include decreased

energy expenditure and reductions of the metabolic rate, and are paralleled by changes in

metabolic hormones. In mammals, starvation induces rapid changes in the concentration of

metabolic hormones, including;

1. Decreased insulin (Lyle et al, 1984; Nair et al, 1987; Webber & Macdonald,

1994; Mercer et al, 1998; Dallman et al, 1999),

Oft Chapter 1 - Literature Review

2. Increased glucagon (Stout et al, 1976; Goschke & Tholen, 1977; Nair et al,

1987),

3. Increased corticosterone (Dallman et al, 1999),

4. Decreased glucose (Stout et al, 1976; Lyle et al, 1984; Nair et al, 1987; Webber

& Macdonald, 1994; Dallman et al, 1999),

5. Decreased triiodothyronine (Harris et al, 1978; Nair et al, 1987; Ikeda et al,

1991; Webber & Macdonald, 1994; Van der Wal et al, 1998),

6. Decreased thyroxine (Harris et al, 1978; Ikeda et al, 1991; Van der Wal et al,

1998),

7. Decreased thyroid stimulating hormone (Harris et al, 1978; Ikeda et al, 1991)

and,

8. Increased growth hormone (Goschke & Tholen, 1977).

Avian species respond to starvation in a similar manner to mammalian species. However,

they differ from mammalian species in that glucagon rather than insulin is the primary

regulator of metabolism, especially during food deprivation (Hazelwood, 1986). In birds

voluntary or imposed starvation induces the following changes in the concentration of

metabolic hormones:

1. Insulin unchanged, or increased (Johnson & Hazelwood, 1982; Rosebrough et

al, 1984; Cherel et al, 1988),

2. Increased glucagon (Cieslak & Hazelwood, 1983; Hazelwood, 1986; Cherel et

al, 1988),

3. Increased corticosterone during phase III of incubation fast, and during imposed

short-term starvation (Cherel et al, 1988; Lea et al, 1992; Geris et al, 1999),

4. Plasma glucose unchanged during incubation or decreased during imposed

starvation (Hazelwood & Lorenz, 1959; Harvey et al, 1978; Le Maho et al,

1981; Rosebrough etal, 1984),

5. Decreased triiodothyronine (Cherel et al, 1988; Groscolas & Leloup, 1989; Lea

et al, 1992; Kiihn et al, 1993; Schew et al, 1996; Darras et al, 1997; Prem et al,

1998; Geris et al, 1999; Van der Geyten et al, 1999),

6. Thyroxine decreased during incubation and either decreased, increased or

unchanged during imposed starvation (Cherel et al, 1988; Groscolas & Leloup,

1989; Kiihn et al, 1993; Schew et al, 1996; Darras et al, 1997; Prem et al, 1998;

Geris et al, 1999; Van der Geyten et al, 1999),

7. Decreased thyroid stimulating hormone (Geris et al, 1999; Van der Geyten et al,

Chapter 1 - Literature Review z'

1999), and

8. Increased or decreased growth hormone (Harvey et al, 1978; Hazelwood, 1986;

Kiihn et al, 1993; Schew et al, 1996; Darras et al, 1997; Geris et al, 1999).

In both avian and mammalian species these changes in metabolic hormones act to maintain

a constant internal environment. In mammals, decreased insulin concentrations decrease

glucose use and promote the use of endogenous substrates for gluconeogenesis (Chilliard et

al, 1998). Hypothyroidism decreases basal metabolism and reduces the rate of protein and

fat turnover (Chilliard et al, 1998). The increased growth hormone concentration has a

sparing effect on protein reserves (Norrelund et al, 2001). These hormonal changes

minimize the loss of energy reserves.

A feature of starvation is the utilisation of adipose, and especially in birds, hepatic reserves

(Kleiber, 1961; Hazelwood, 1986). The catabolism of these reserves releases substrates

into the circulation that also alter metabolism. In mammalian species, acute starvation is

associated with an increase in the circulating concentration of free fatty acids (Stout et al,

1976; Goschke & Tholen, 1977; Lyle et al, 1984; Romijn et al, 1991). In avian species a

similar situation is observed during the first few days of fasting or incubation anorexia (Le

M a h o et al, 1981; Hazelwood, 1986). However, with prolonged incubation anorexia the

concentration of free fatty acids decreases (Le M a h o et al, 1981; Groscolas & Leloup,

1989). In humans, there is evidence that the elevation in free fatty acids in response to

starvation suppresses gluconeogenesis and minimises protein degradation (Fery et al,

1996). As such, the response of animals to starvation provides insights into the control of

appetite and the range of energy sparing mechanisms that are employed, when the animal's

response are not influenced by the reproductive state.

Metabolite concentrations also alter in response to imposed or naturally occurring

starvation. The changes in metabolite concentrations can reveal the energy source that the

animal is using to meet its energetic requirements. In avian species, B-hydroxybutyrate

rises rapidly in response to fasting, but at a slower rate in response to naturally occurring

anorexia during incubation (Le M a h o et al, 1981; Cherel et al, 1988). Increases in the

concentration of B-hydroxybutyrate indicate catabolism of adipose stores. The later stages

of incubation anorexia are associated with increased uric acid and decreased B-

hydroxybutyrate plasma concentrations (Le M a h o et al, 1981; Cherel et al, 1988; Groscolas

& Leloup, 1989). This indicates that the birds are catabolising protein and obtaining less

Chapter 1 - Literature Review z o

energy from adipose reserves (Groscolas & Leloup, 1989). It is likely that the emu

undergoes similar changes in metabolites, hormones and energy sparing mechanisms in

response to starvation. As such, investigation of the emu's response to starvation could

provide insights into the control of appetite and the nature of energy sparing mechanisms

employed by the emu.

7.0 - Conclusions

The decreased appetite and diminished adipose reserves that are associated with the

breeding season limit the commercial productivity of the emu. To overcome production

constraints associated with reproduction in the emu it is essential that the physiology and

biochemistry underpinning appetite in the emu be better understood. The information

given in the preceding sections relates to a general integrated model for the control of

energy balance that is based mainly on data from mammalian species. However, the emu

may not adhere to this model given the strong evolutionary pressure applied to it for

efficient use of available energy sources by the harshness of its environment.

The general aim of this study has been to improve the understanding of the control of

appetite and adiposity in the emu. To address this aim, the general hypothesis of this thesis

was that seasonal variation in appetite and adiposity in the emu operates under the general

model developed in other species.

The general hypothesis was tested using experimental models associated with altered

appetite, adiposity and metabolic hormone concentrations. The experimental models used

were high fat diets and dexamethasone treatment (increased appetite and adiposity),

incubation (decreased appetite and adiposity) and starvation (increased appetite and

decreased adiposity). Three approaches were taken to test the general hypothesis. First, to

determine the role of leptin in the control of appetite and adiposity attempts were made to

clone and sequence the e m u leptin gene. Once sequenced it was intended that the gene

would be used to develop an emu specific leptin radioimmunoassay, and investigate

changes in leptin gene expression during the annual cycle of the emu. Second, the

concentration of metabolic hormones was measured in all of the experiments. Finally, to

determine the role of N P Y in the control of appetite and adiposity the m R N A expression of

N P Y from the mediobasal hypothalamus and preoptic area was quantified in the

glucocorticoid treatment, incubation and starvation experiments.

Chapter 2

General Materials and Methods

Page

1. Experimental location 30

2. Animals 30

3. Nutrition 30

4. Measurements 30

5. Blood sampling 33

6. Euthanasia 34

7. Tissue collection techniques 34

8. Metabolic substrate and hormone assays 36

9. Hormone data analysis 49

10. Molecular techniques 52

Chapter 2 - General Materials and Methods 30

A general overview of the materials and methods employed during the course of the

study is presented here. Details of the techniques, treatments and analyses will be given

fully in the experimental chapters. All experiments were approved by the University of

Western Australia animal ethics committee and were carried out in accordance with

welfare guidelines. Appendix 2 contains a list of all suppliers mentioned in this and

subsequent chapters.

1. Experimental Location

Experiments were performed at the Shenton Park research station (31°56'S), part of the

University of Western Australia, situated in Perth, Western Australia. Perth has a

mediterranean climate, with wet winters (average rainfall 150.9 m m ; daily temperature

range 8.1 -28.3°C) and hot, dry summers (average rainfall 12 m m ; daily temperature

range 14.8-46.7°C) (Bureau of Meterology website). All animals used in the

experiments were maintained outdoors in individual pens (pen size: experiments 1, 3

and 4 were greater than 15 m2, experiment 2 were approximately 4 m ) .

2. Animals

All emus used during the course of this research were either bred at the Shenton Park

facility ( U W A ) , Medina (Agriculture Western Australia) or obtained from a commercial

breeder in the vicinity of Perth (P & T Amrein, Wungong). The animals were

acclimatised at the Shenton Park facility in individual pens for at least two weeks before

the commencement of any experiments. The animals used were sexually mature adults,

ranging in age from 14 months to five years old, unless otherwise stated. Due to a

downturn in the emu market birds were not readily available and therefore the choice of

sex for each experiment was determined by availability.

3. Nutrition

Animals were fed an emu breeder pellet obtained from Glen Forrest Stockfeeders (for

specifications see Appendix 2). Feed and water were provided ad libitum for all

experiments, unless otherwise stated.

4. Measurements

Live weight


Animals were weighed periodically during the course of all the experiments. The birds

were caught, manoeuvred onto livestock scales (Ruddweigh K M - 2 electronic weighing

system), and held while recordings were made (Fig. 2.1).

Figure 2.1: Live weight measurement of the emu.

Fat score

A n assessment of the size of the subcutaneous fat depot, as developed by Mincham et al

(1998), was conducted periodically for experiment 2. This entailed restraining the bird

while the region between the preacetabular ilium and postacetabular ilium was palpated.

The body condition of the animal in terms of build up of muscle and fat around the

backbone was scored between 1 and 5. The values correspond to the following

descriptions:

1 - bones prominent and sharp;

2 - bones prominent but smooth;

3 - bones can be felt but are smooth and rounded;

4 - bones are detectable with pressure on the thumb;


Feed Intake

Throughout the experiments individual feed intakes were recorded. Buckets were filled

with feed such that the combined weight was in the range of 6 to 8 kilograms. The

exact weight was recorded and the buckets returned to the pens. After periods of either

48 or 72 hours the buckets were collected and the combined weight of the bucket and

the remaining feed was recorded. The buckets were then refilled, weighed, recorded

and returned to the bird. Weighing and refilling of buckets was done in the morning.

Blood glucose

Blood glucose was measured during experiments' 2 and 3, using a hand-held blood

glucose meter (Precision Q.I.D, Medisense) by applying a droplet of blood to an

electrode (Precision plus, Medisense) immediately following each bleed. The blood

glucose levels recorded for emus ranged from 5.1 to 16.5 mmol/L. The between

electrode variation determined by the manufacturer is 2.1-3.7% over the range of

concentrations observed for emus. Parallelism had been previously determined for

emus (Van Cleeff, 2001).

Carcass composition

Following the removal of tissue samples (further details are given in the tissue

collection techniques below), the carcass was stripped of subcutaneous adipose tissue.

Following this the internal organs were removed and the visceral fat dissected out. The

tissues of interest were then weighed with the whole procedure taking approximately 45

minutes per bird (Table 2.1). The digestive tract length (proventriculus to cloaca) was

also measured for experiment 1 (Chapter 4).

Table 2.1: Tissues weighed for each experiment

Tissue

Subcutaneous adipose

Visceral adipose

Liver

Gastrocnemius lateralis (hamstring)

Ovary

Testis

Heart

Digestive tract (proventriculus to cloaca)

1

/

•/

Experiment

2 3

V

s

s s s V

4

Experiments 1, 2, 3 and 4 are described in Chapters 4,5, 6 and 7 respectively


Tissue fat content determination

Following slaughter the fat content of liver and muscle was determined for experiment 1

using the method developed within the Animal Science group and described by Pluske

(1993). Approximately 2 g (wet weight) samples of each tissue were collected in

triplicate. Samples were frozen at -20°C, freeze-dried and re-weighed. The tissues were

then ground to a powder and mixed with 40 ml of solvent (hexanedsopropanol = 3:2),

and tumbled at a slow speed for two hours before being left to stand overnight. The

samples were filtered under vacuum through Whatman no. 1 filter paper over a ceramic

filter with 3 micron pores. The filtrate was dried for two hours at 70°C to evaporate

excess solvent and weighed. The fat content was determined as the amount removed by

the solvent:

fat content = freeze-dried weight - weight of tissue after solvent extraction.

Figure 2.2: Jugular bleeding of the emu.

5. Blood sampling

The animals were caught and restrained by holding their wings while standing behind

them. This was necessary to control the animal while being bled. Blood samples were

taken at regular intervals from the jugular using an 18-gauge needle and either a 10 or

20 ml syringe (Fig. 2.2). Between 10-20 ml samples were collected and immediately

transferred into plastic tubes for plasma or serum collection. Plasma collection tubes

contained 40 IU/ml of heparin with a small amount of polystyrene granules (Kwikspin

granules: Disposable Products Pty Ltd, Australia) added. Plasma tubes were centrifuged


(Sarstedt Australia Pty Ltd) and were stored at 4°C for 1-3 days before being

centrifuged. All centrifugation of blood samples was performed at 3000 rpm and 4°C

for 25 minutes. Plasma and serum samples were decanted into plastic tubes (Sarstedt

Australia Pty Ltd) and stored at -20°C. All blood samples were collected in the

morning.

6. Euthanasia:

All birds were euthanased at the completion of experiments by the injection of 25 ml of

Lethabarb (sodium pentabarbitone 325 mg/ml, Virbac) into the jugular.

7. Tissue collection techniques for:

RNA extraction

Samples were collected for R N A extraction from the following tissues: adipose, liver,

mediobasal hypothalamus and preoptic area. To obtain the mediobasal hypothalamus

and preoptic area the head was removed following euthanasia and the skull opened

using secateurs to cut through the skull in a hexagonal pattern (Fig. 2.3). The piece of

skull bounded by the cuts was then pried up along the incision at the back of the skull,

and the dura mere carefully cut away to remove it without disturbing the brain.

Incisions were made through the cerebellum and the optic chiasma, and the brain gently

lifted free by cutting through any constraining connective tissue and nerves. The region

of the brain containing the mediobasal hypothalamus and the preoptic area were

collected and snap frozen in liquid nitrogen for R N A extraction (Fig. 2.4). All tissues

required were excised within 15 minutes of the animal being euthanased and cut into

appropriately sized pieces (not greater than one gram). To prevent R N A degradation by

RNases prior to tissue collection all instruments used were wrapped in alfoil and baked

at greater than 200°C for 24 hours, and any manipulation of tissues was carried out

either on baked alfoil or in baked glass petri dishes. Where instruments were re-used

they were rinsed in a 2 % S D S solution between animals. Sterile 5 ml tubes (Sarstedt

Australia Pty Ltd) were used to store tissues and samples were snap frozen in liquid

nitrogen as rapidly as possible after collection (less than one minute). Samples were

stored at -80°C until R N A extractions were performed (section 10.).


Figure 2.3: An emu skull showing the placement of incisions through the skull for brain removal.

Figure 2.4: Schematic saggital view of the avian brain showing the microdissection sites for collection of the mediobasal hypothalamus ( M B H ) and preoptic area (POA) for R N A extraction and R P A (CER=cerebellum, PS=pituitary stalk, OC=optic chiasma). Adapted from Pearson (1972).


8. Metabolic substrate and hormone assays

General buffers and reagents

0.5 M phosphate buffer To -800 ml of nanopure water add 61.3 g disodium

hydrogen orthophosphate (Na2HP04) and 10 g

sodium dihydrogen orthophosphate

(NaH2PO4.2H20), adjust the p H to 7.5 and bring the

volume to 1 L with nanopure water,.

0.3 Mphosphate buffer 3:2 dilution of 0.5 M phosphate buffer in nanopure

water, p H 7.5.

0.1 Mphosphate buffer 1:4 dilution of 0.5 M phosphate buffer in nanopure

water, p H 7.5.

Phosphate buffered saline (PBS) 100 ml 0.1 M phosphate buffer, 8.9 g sodium

chloride (NaCL), 1 g sodium azide (NaN3) make to 1

L with nanopure water, p H 7.5.

Sample preparation

E m u serum and plasma often exhibit high lipid contents. The lipid interferes with the

detection of luteinising hormone (LH), glucagon and insulin via radioimmunoassay. To

remove the lipid two approaches were employed. Samples to be assayed for L H and

glucagon (1 ml) were treated with 4 5 % polyethylene glycol (PEG) (200 /il), incubated

for 30 minutes and centrifuged at 1600 g for 10 minutes (Blache et al, 2001a). The

clear plasma or serum was transferred to a fresh tube. P E G extraction recovers 75%,

8 9 % and 4 0 % respectively of LH, glucagon and insulin (Van Cleeff, 2001). As the

recovery of insulin by P E G extraction is low, lipid removal from samples to be assayed

for insulin was achieved by centrifugation at high speed for between 10 and 30 minutes.

The clear plasma or serum was removed from below the fat layer to a fresh tube.

Corticosterone

Corticosterone was measured using extraction methods developed by Smith et al (1972)

and Etches (1976).

Buffers/reagents

Gelatin phosphate buffer (GPB) PBS (pH 7.5) + 1 g gelatin/L

Dextran-coated charcoal 6.25 g charcoal + 0.625 g dextran T70/L in GPB.


Standards

A stock solution of corticosterone (4-pregnene-llp, 21-diol-3, 20-dione, Sigma) was

prepared in ethanol to a concentration of 7.6 jtfg/ml. Standards were made by serial

dilution to the following concentrations: 20,10, 5, 2.5,1.25,0.6125, 0.306, 0.15 and

0.078 ng/jiil in ethanol.

Antiserum

The first antibody, corticosterone antiserum B3-163, was raised in rabbits and obtained

from Endocrine Sciences Products. To use in the assay it was reconstituted, and diluted

to 1:200 in gelatin phosphate buffer.

Tracer

[1,2,6,7-3H] corticosterone (Amersham Biosciences Pty Ltd).

Purification:

A 5 ml pipette stoppered with a glass bead was filled with 2.5 ml of LH-20 sephadex

(Pharmacia Biotech) swollen in a mixture of benzene:methanol (9:1). 10 /xCi of tracer

was dried down and redissolved in benzene:methanol (9:1) (100 (il). This was added to

the column and allowed to enter the gel bed. The column was eluted with

benzene:methanol (9:1). Fractions (500 ptl) were collected into glass tubes and an

aliquot of 5 (i\ of each fraction was counted with scintillint (Starscint, Packard) using a

liquid scintillation counter (Packard 1500 Tri-Carb Liquid Scintillation Analyzer). The

appropriate fractions were pooled, dried down under compressed air and dissolved in

absolute ethanol (5 ml). O n Day 1 of the assay the tracer was diluted to 10000 cpm per

20 ul

Assay procedure

The standard curve was placed at the beginning of the assay. Each standard curve

included 3 tubes for total counts (TC), 3 tubes for non-specific binding (NSB), 9

replicates of zero binding (B0), 3 replicates of each standard and 4 replicates each of two

pool samples.

On Day 1, the tubes to be used for the assay were coated with 10000 cpm of the tracer

prior to commencement of the assay. 12 x 75 m m glass tubes were coated for samples

and pools and 10 x 75 m m glass tubes were coated for standards, TC, N S B and Bo.

Tracer (20 /i\, 10000 cpm) was added to all tubes, standards (100 /xl) were added in


triplicate to the appropriate tubes. All tubes were evaporated to dryness under

compressed air at 37°C using a heated block.

Extraction:

Samples of unknown serum and pools (100 p\) were added to the coated tubes, vortexed

individually and incubated for 30 minutes at room temperature. The samples and pools

were extracted by adding 2 ml of 2,2,4-trimethyl pentane (iso-octane) or hexane to each

tube and vortexing twice for 30 seconds. The tubes were then centrifuged at 1500 rpm

for 5 minutes (Beckman J-6M/E centrifuge). The aqueous layer was frozen by partially

immersing the tubes in an acetone-dry ice bath, the solvent layer was then discarded.

Dichloromethane (2 ml) was added to each tube and the tubes were vortexed twice for

30 seconds. The tubes were centrifuged at 1500 rpm for 5 minutes (Beckman J-6M/E

centrifuge). The tubes were returned to the acetone-dry ice bath to freeze the upper

aqueous layer, the organic layer was then poured off into clean 10 x 75 m m glass tubes.

The dichloromethane was evaporated to dryness under compressed air at 37°C. First

antibody (250 /xl) was added to all tubes except the T C and N S B (GPB (250 fil) was

added to T C and N S B in place of the first antibody). The tubes were then shaken and

incubated for 24 hrs at 4°C.

On Day 2, dextran-coated charcoal (200 ^tl) was added to all tubes except TC the tubes

were incubated for 20 minutes at 4°C before being centrifuged at 1500 rpm for 10

minutes (Beckman J-6M/E centrifuge). A volume of 400 fx\ of the supernatant was then

removed and added to a counting vial along with 2 ml of scintillant (Starscint, Packard)

and counted for 3 minutes each in a liquid scintillation counter (Packard 1500 Tri-Carb

Liquid Scintillation Analyzer).

Glucagon

Glucagon was measured using Linco kit tracer, standards and first antibody (Linco

Research, Inc.). The assay was validated for emus by Van Cleeff (2001).

Buffers/reagents

Glycine buffer 15 g glycine, 10 g bovine serum albumin (BSA), 9

g ethylenediaminetetra acetic acid (EDTA) (non-

salt), 1 g NaN3. Dissolve in 800 ml of nanopure


water, adjust p H to 8.8 and bring volume to 1 L

with nanopure water.

Tracer buffer 0.375 g glycine, 2.5 g BSA, 2.25 g E D T A (non-

salt), 0.2 g NaN3. Dissolve in 150 ml of nanopure

water, adjust p H to 8.8 and bring volume to 250 ml


Standards

Purified recombinant human glucagon (800 pg/ml) was used to make the standards by

serial dilution to the following concentrations: 400, 200, 100, 50, 25 and 12.5 pg/ml in

glycine buffer.

Antiserum

First antibody:

Guinea pig anti-glucagon, specific for pancreatic glucagon. Reconstituted on arrival

with 100 ml of glycine buffer and 100 fxg of aprotinin.

Second antibody:

Sheep anti-guinea pig raised in the laboratory and diluted 1:20 in glycine buffer with

normal guinea pig serum (NGPS) added at a dilution of 1:400.

Tracer

125I-glucagon obtained from Linco. Reconstituted by adding 27 ml of tracer buffer per

vial, and occasional gentle mixing for 30 minutes. This was further diluted to >12,000

counts per 100 id.

Assay procedure


included 3 tubes for TC, 3 tubes for NSB, 9 replicates of Bo, 3 replicates of each

standard and 4 replicates each of three pool samples.

On Day 1, unknown plasma samples (50 fi\), and pools and standards (100 /*1 each)

were diluted to a final volume of 200 y\ with glycine buffer in 12 x 75 m m plastic tubes

(Sarstedt Australia). Glycine buffer was added to N S B (300 id) and B 0 (200 /d) tubes.

The first antibody preparation (100 id) was added to all tubes except the N S B and TC.

Tubes were then incubated overnight at 4°C.


O n Day 2, tracer (100 ul) was added to all tubes and mixed. The tubes were incubated

overnight at 4°C.

On Day 3, second antibody (100 /d) was added to all tubes except the TC, mixed and

incubated overnight at 4°C.

On Day 4, 3% PEG in PBS (pH 8.8) (1 ml) was added to all tubes except the TC, before

centrifugation at 3000 rpm for 25 minutes at 4°C (Beckman J-6M/E centrifuge) after

which the supernatant was poured off. The T C were not centrifuged or poured off. The

activity of the precipitate was counted for 90 seconds each in a gamma counter (Packard

Cobra II Auto-Gamma).

Insulin

Insulin was measured in plasma or serum using the method for chickens developed by

McMurtry et al (1983). This method was validated for emu plasma and serum by Van

Cleeff (2001). The chicken insulin used for the standards and tracer, and the first

antibody were obtained from Professor McMurtry, U S D A.

Buffers/reagents

Insulin buffer 1.095 g NaH2P04.2H20, 5.906 g Na2HP0 4,

18.6125 g E D T A (disodium salt). Dissolve in 800

ml of nanopure water, adjust p H to 7.5, and bring

volume to 1 L.

Chloramine-T0.3 Mphosphate 5 m g chloramine-T in 10 ml 0.3 M phosphate

buffer, prepared 10 minutes before the iodination.

Sodium metabisuphite 5 m g sodium metabisulphite (Na2S205) in 10 ml

0.3 M phosphate buffer.

Potassium iodide 200 m g potassium iodide (KI) in 10 ml 0.1 M

phosphate buffer.

Standards

Standards were made by serial dilution of a 10 ng/ml stock of chicken insulin to the

following concentrations: 7.5, 5, 3.75, 2.5, 1.875, 1.25, 0.9375, 0.625,0.46875, 0.3125,

0.234 and 0.15625 ng/ml in insulin buffer + 0.1% gelatine.


Antiserum

First antibody:

The first antibody was raised in guinea pigs against chicken insulin. It was used at

1:15,000 with 1:600 of normal guinea pig serum diluted in insulin buffer + 0.1%

gelatine.

Second antibody:

Sheep anti-guinea pig, raised in the laboratory and diluted 1:10 with insulin buffer.

Tracer

Columns:

G100 (Pharmacia) equilibrated with 1 % BSA, 0.1 M phosphate buffer

Iodination:

Chicken insulin was stored as 5 iig aliquots at -80°C. To a 5 fig aliquot, 0.5 mCi of I

and chloramine-T buffer (10 id) was added, mixed and allowed to incubate at room

temperature for 90 seconds. Potassium iodide (100 pA) and sodium metabisulphite (100

/d) solutions were added to stop the reaction. The reaction was added to the column,

and the reaction vessel rinsed with 1 % BSA, 0.1 M phosphate buffer (100 itl). The

rinse solution was also added to the column and 1 ml fractions were eluted with 1 %

BSA, 0.1 M phosphate buffer. A n aliquot of 10 /d of each fraction was counted for 10

seconds in a gamma counter (Packard Cobra II Auto-Gamma) to identify the appropriate

fractions. The tracer was immediately diluted to >15,000 counts/100 itl with insulin

buffer + 1 % B S A and added to the assay. Excess tracer was stored at 4°C.

Assay procedure


included 3 tubes for TC, 4 tubes for NSB, 9 replicates of Bo, 3 replicates of each

standard and 4 replicates each of two pool samples.

On Day 1, unknown plasma samples and pools (100 itl) were diluted to 200 /d with

insulin buffer + 0.1% B S A (100 p\) in 12 x 75 m m plastic tubes (Sarstedt Australia).

Standards (200 fi\) were added to appropriate tubes. Insulin buffer + 0.1% B S A was

added to N S B (300 /d) and B 0 (200 iil). The first antibody preparation (100 itl) was

added to all tubes except the N S B and TC. Tubes were then incubated overnight at 4°C.


O n Day 2, tracer (100 itl) was added to all tubes and mixed. The tubes were incubated

for 24 hours at 4°C.

On Day 3, second antibody (100 fi\) was added to all tubes except the TC, mixed and

incubated overnight at 4°C.

On Day 4, 6% PEG in PBS (1 ml) was added to all tubes except the TC, before

centrifugation at 3000 rpm for 25 minutes at 4°C (Beckman J-6M/E centrifuge) after

which the supernatant was poured off. The T C were not centrifuged or poured off. The

activity of the precipitate was counted for 1 minute each in a gamma counter (Packard

Cobra II Auto-Gamma).

Luteinising hormone (LH)

L H was measured in plasma using the method described by Talbot et al (1988), based

on the original L H assay developed by Follet et al (1972). The validation of this method

for emu plasma is described by Malecki et al (1998) and Blache et al (2001a). The

chicken L H fraction (PRC-AEI-s-1) used for the assay standards and iodination was

kindly donated by Professor Sharp, Roslin Institute, Midlothian, Scotland. This was

prepared using the method described by Talbot et al (1988). The fraction was purified

by passage through a G100 column (Pharmacia Biotech) and had a potency of 0.43

(0.375-0.499) NIH-0-LH-S19 in the chicken granulosa cell assay (Hammond et al,

1980).

Buffers/reagents

LH buffer 80 ml 0.5 M phosphate buffer, 8.75 g NaCl, 10

m M EDTA, 1 g NaN3, 1 g BSA, bring to 900 ml

with nanopure water, adjust p H to 7 with 1 N

sodium hydroxide (NaOH), and bring to 1 L with

nanopure water.

0.1 Mphosphate-gelatine 0.1 M phosphate buffer, 0.25% gelatine in

nanopure water, p H 7.5.

0.1 Mphosphate-BSA 0.1 M phosphate buffer, 1 % BSA.

Chloramine-T 0.1 Mphosphate 5mg chloramine-T in 10 ml 0.1 M phosphate

buffer, prepared 10 minutes before the iodination.

Sodium metabisuphite 10 m g Na2S20s in 100 ml 0.1 M phosphate buffer.


Potassium iodide 1 g KI in 10 ml 0.1 M phosphate buffer.

Standards

Aliquots of 1.25 (xg of L H in nanopure water were stored at -20°C. The L H solution

was diluted with L H buffer to give a 250 ng/ml working stock solution. Standards were

made by serial dilution to the following concentrations: 10, 5, 2.5, 1.25, 0.625, 0.312,

0.156, 0.078, 0.039 and 0.019 ng/ml in L H buffer.

Antiserum

First antibody:

Raised in a rabbit immunised against the chicken L H preparation supplied by Professor

Sharp. The stock (1:200 dilution) was diluted further with L H buffer to give a working

dilution of 1:20,000.

Second antibody:

Donkey anti-rabbit raised in the laboratory. The stock was diluted with L H buffer (1:6)

to give the working solution.

Tracer

Columns:

G25 column (Pharmacia) equilibrated with 0.1 M phosphate-gelatine buffer + 2 % BSA.

Iodination:

To 1.25 [ig L H in 10 ill nanopure water, 0.5 mCi Na-125I and chloramine-T buffer (20

itl) were added and incubated for 60 seconds. The reaction was stopped by adding

sodium metabisulphite (100 p.1) and potassium iodide (200 id). The column was

drained down to the gel surface. The contents of the reaction vessel were then placed on

the column. The reaction vessel was rinsed with potassium iodide buffer (250 /d) that

was then added to the column. Fractions (500 /d) were eluted with 0.1 M phosphate-

gelatine buffer into tubes containing 500 ill 0.1 M phosphate-BSA buffer. O n Day 2 of

the assay the appropriate fraction was diluted to 10,000 counts per 50 ill with L H buffer.

Assay procedure


included 3 tubes for TC, 3 tubes for N S B , 9 replicates of Bo, 3 replicates of each

standard and 4 replicates each of two pool samples.


O n Day 1, unknown plasma samples and pools (50 ul) were diluted to 200 ill with L H

buffer in 12 x 75 m m plastic tubes (Sarstedt Australia). Standards (200 til) were added

to the appropriate tubes. L H buffer was added to N S B (250 ul) and B 0 (200 ul) tubes.

The first antibody (50 ul) was added to all tubes except the N S B and TC. Tubes were

then incubated overnight at 4°C.

On Day 2, tracer (50 /d) was added to all tubes and vortexed briefly. The tubes were

incubated for 48 hours at 4°C.

On Day 4, NRS (1:400 dilution with LH buffer) (50 ul) and second antibody (50 ul)

were added to all tubes except the TC, vortexed briefly and incubated overnight at 4°C.

On Day 5,6% PEG in LH buffer (1 ml) was added to all tubes except the TC, before

centrifugation at 1500 g for 30 minutes at 4°C (Beckman J-6M/E centrifuge) after which

the supernatant was poured off. The T C were not centrifuged or poured off. The

activity of the precipitate was then counted for 1 minute each in a gamma counter

(Packard Cobra II Auto-Gamma).

Prolactin

Prolactin was measured in plasma using the method developed by Talbot and Sharp

(1994). Validation of this method for emu prolactin is described by Malecki et al

(1998). Recombinant derived chicken prolactin used for the assay standards, raising the

first antibody and iodination was kindly donated by Professor Sharp, Roslin Institute,

Midlothian, Scotland. Recombinant chicken prolactin was prepared using the method

described by Hanks et al (1989).

Buffers/reagents

Chloramine-T0.3 Mphosphate 5 m g chloramine-T in 10 ml 0.3 M phosphate

buffer.

2 % BSA 0.2 g B S A in 10 ml 0.3 M phosphate buffer.

Tris/Tween 20 (make fresh) 1.8 g NaCl, 3.14 g tris/HCl, 0.1 g NaN3,200 id

Tween 20, bring to 200 m L in nanopure water,

adjust p H to 8.

Prolactin buffer 900 ml 0.1 M phosphate buffer (pH7.5), 9 g NaCl,

3.72 g E D T A (disodium salt), 1 g NaN3,1 g BSA.


Adjust p H to 7 with 1 N N a O H and bring to 1 L


Standards

The prolactin stock was diluted with prolactin buffer to give a 1250 ng/ml working

stock solution. Standards were made by serial dilution to the following concentrations:

125, 62.5, 31.25, 15.625, 7.8125, 3.9, 1.95, 0.977, 0.4883 and 0.24 ng/ml in prolactin

buffer.

Antiserum

First antibody:

Raised in a rabbit immunised against recombinant chicken prolactin by Professor Sharp.

The stock (1:100 dilution) was diluted further with prolactin buffer to give a 1:6000

working solution.

Second antibody:

Donkey anti-rabbit raised in the laboratory. The stock solution (1:2 dilution) was

further diluted with prolactin buffer (1:1) to give the working solution.

Tracer

Columns:

PD10 (Pharmacia Biotech) and G100 (Pharmacia Biotech) equilibrated with Tris/Tween

20 buffer.

Iodination:

Prolactin stock (14 ul, 5 fig) was aliquoted into an eppendorf tube and 0.3 M phosphate

buffer (16 ul), 125I-Na (5 ul) and chloramine-T buffer (10 til) were added. The reaction

vessel was vortexed and incubated at room temperature for 60 seconds. The reaction

was stopped by the addition of 0.3 M phosphate buffer (200 id). The reaction solution

was applied to a P D 10 column and the reaction tube rinsed with 0.3 M phosphate

buffer (250 ill) the rinse product was also applied to the column. Fresh Tris/Tween 20

buffer was used to elute 11-drop fractions. The appropriate fractions were identified by

counting 10 id aliquots using a gamma counter (Packard Cobra II Auto-Gamma), and

then combined. The G 100 column was allowed to drain down to the gel surface and the

iodinated fraction applied. Fractions (1 ml) were eluted with Tris/Tween 20 buffer. The

appropriate fraction was identified by counting 10 id aliquots using a gamma counter


(Packard Cobra II Auto-Gamma). The fraction was diluted 1:1 with 2 % BSA. O n Day

2, the appropriate fraction was diluted to 12,000 counts per 50 /d with prolactin buffer.

Assay procedure


included 3 tubes for TC, 3 tubes for NSB, 9 replicates of B0, 3 replicates of each


On Day 1, unknown plasma samples and pools (100 id) were added to 12 x 75 mm

plastic tubes (Sarstedt Australia). The standards (100 ul) were added to appropriate

tubes. The first antibody (50 ul) was added to all tubes except the N S B and TC.

Prolactin buffer (150 id and 100 id respectively) was added to N S B and B 0 tubes.

Tubes were incubated overnight at 4°C.

On Day 2, tracer (> 12000 counts/50 ul) (50 ul) was added to all tubes and vortexed

briefly. The tubes were incubated for 24 hours at 4*C.

On Day 3, NRS (1:600 dilution with prolactin buffer) (50 ul) and second antibody (50

ul) were added to all tubes except the TC, vortexed briefly and incubated overnight at

4°C.

On Day 4, 6% PEG in PBS (1 ml) was added to all tubes except the TC, before

centrifugation at 1500 g for 30 minutes at 4°C (Beckman J-6M/E centrifuge) after which

the supernatant was poured off. The T C were not centrifuged or poured off. The

activity of the precipitate was then counted for 1 minute each in a gamma counter

(Packard Cobra II Auto-Gamma).

Thyroid Hormones

Total triiodothyronine (T3) and total thyroxine (T4) levels in serum and plasma were

measured using a non-extraction assay described by Dawson et al (1996) and Dawson

and Deeming (1997). The validation of this method for emu plasma is described by

Blache et al (2001c).


Buffers/reagents

Barbitol buffer 12 A g barbital sodium (C8HiiN2Na03) make up to

1 L in nanopure water, p H 8.6.

ANSA buffer 0.8 g 8-anilino-l-naphthalene sulphonic acid

(ANSA) to 100 ml of barbitol buffer.

T3 standards

A stock solution of T3 was prepared by dissolving 10 m g of T3 powder in 100 ml of

barbitol buffer. It was further diluted 1:1000 to give a final concentration of 100 ng/ml.

Standards were made by serial dilution to the following concentrations: 50, 25,12.5,

6.25, 3.125, 1.56, 0.78, 0.39, 0.195, 0.0975, 0.049, 0.0245, 0.012 and 0.006 ng/ml in

barbitol buffer.

T3 antiserum

First antibody:

Rabbit polyclonal antibody anti-T3 was purchased from Biogenesis Immuno

Diagnostics, and diluted 1:10 on arrival in barbitol buffer. For the assay it was diluted

1:40 to give a 1:400 stock. The first antibody was added to the assay at a working

dilution of 1:12,000.

Second antibody:

Donkey anti-rabbit raised in the laboratory and diluted 1:5 with barbitol buffer.

T3 tracer

L-3,5,3'-[125I]- triiodothyronine (NEN Life Science Products, Inc.) diluted in A N S A

buffer to give 10,000 cpm/20 ul

T3 assay procedure


included 3 tubes for TC, 3 tubes for N S B , 9 replicates of Bo, 3 replicates of each


O n Day 1, standards (20 id) were diluted to a total volume of 60 ul, with charcoal

stripped plasma (20 ul) and barbitol buffer (20 ul). The unknown serum samples and

pools (20 ul) were diluted to a total volume of 60 ul with barbitol buffer (40 ul). 10 x

75 m m glass tubes were used (Sarstedt Australia). The first antibody (20 id) was added


to all tubes except the N S B and TC. Tracer (20 ul) was added to all tubes and vortexed

briefly. The tubes were incubated for 48 hours at 4°C.

On Day 3, second antibody and normal rabbit serum (NRS) (1:500 dilution) (20 ul each)

were added to all tubes except the TC, and vortexed briefly. The tubes were then

incubated for 24 hours at 4°C.

On Day 4, 6% PEG in barbitol buffer (1 ml) was added to all tubes except the TC,

before centrifugation at 1500 g for 25 minutes at 4°C (Beckman J-6M/E centrifuge)

after which the supernatant was aspirated. The T C were not centrifuged or aspirated.

The activity of the precipitate was then counted for 1.5 minutes each in a gamma

counter (Packard Cobra II Auto-Gamma).

T4 standards

A working stock solution of T4 was made by adding 100 ul of a 12.8 umol/L

concentrated stock solution to 10 ml of barbitol buffer. Standards were made by serial

dilution to the following concentrations: 64, 32, 16, 8,4, 2,1,0.5, 0.25 and 0.125 nM/L

in barbitol buffer.

T4 antiserum

First antibody:

Rabbit polyclonal antibody anti-T4 was purchased from Biogenesis Immuno

Diagnostics, and diluted 1:10 on arrival in barbitol buffer. For the assay it was diluted

1:40 to give a 1:400 stock. The first antibody was added to the assay at a working

dilution of 1 in 6,400.

Second antibody:

Donkey anti rabbit raised in the laboratory and diluted 1:5 with barbitol buffer.

T4 tracer

L-[125I]-thyroxine (NEN Life Science Products, Inc.) diluted in A N S A buffer to give

10,000 cpm/20 id.

T4 assay procedure


included 3 tubes for TC, 3 tubes for NSB, 9 replicates of B0, 3 replicates of each



O n Day 1, standards (20 ul) were diluted to a total volume of 60 ul, with charcoal

stripped plasma and barbitol buffer (20 ul each). The unknown serum samples and

pools (20 ul) were diluted to a total volume of 60 ul with barbitol buffer (40 ul). 10 x

75 m m glass tubes were used (Sarstedt Australia). The first antibody (20 ul) was added

to all tubes except the N S B and TC. Tracer (20 ul) was added to all tubes and vortexed

briefly. The tubes were incubated for 24 hours at 4°C.

On Day 2, second antibody and NRS (1:500 dilution) (20 ul each) were added to all

tubes except the TC, and vortexed briefly. The tubes were then incubated for 24 hours

at 4°C.

On Day 3, 6% PEG in barbitol buffer (1 ml) was added to all tubes except the TC,

before centrifugation at 1500 g for 25 minutes at 4°C (Beckman J-6M/E centrifuge)

after which the supernatant was aspirated. The T C were not centrifuged or aspirated.

The activity of the precipitate was then counted for 1.5 minutes each in a gamma

counter (Packard Cobra II Auto-Gamma).

9. Hormone data analysis

Calculation of hormone assay results

The final counts obtained for each assay were saved onto M S - D O S disks and converted

into Macintosh format. Concentrations of the hormones were then calculated using the

AssayZap Universal Assay Calculator.

Sensitivity and precision

The N S B , Bo, and assay sensitivity for each hormone assay are summarised in Table 2.2.

The sensitivity of the assay is defined as the limit of detection of the standard curve.

The precision of the assay expressed as intra-assay and inter-assay coefficients of

variation are given in Table 2.3. For each hormone assayed, the intra-assay coefficient

of variation was calculated from the standard deviation among replicates of each quality

control sample. In Tables 2.2 and 2.3, the assay statistics and intra-assay coefficients of

variation for glucagon, T3, T4 and corticosterone are presented separately for the assays

performed, because the hormone concentrations from the samples were measured within

one assay for each experiment.

Chapter 2 - General Materials and Methods

Pools for assessing intra-assay coefficients of variation

Whenever possible, plasma pools were prepared for high, medium and low

concentrations of each hormone.


Table 2.2: Assay statistics.

Assay

PRL ng/ml (1) 1 (exp 3) T3 ng/ml (3) * 1 (exp 1) 2(exp 2) 3 (exp 3 & 4) T4 nM/L (3) * 1 (exp 1) 2(exp 2) 3 (exp 3 & 4) C O R T ng/jtl (4) * 1 (exp 2A) 2 (exp 2B) 3 (exp 3 & 4) 4 (exp 3 & 4, repeats) INS ng/ml (3) * 1 (exp 1) 2 (exp 2) 3 (exp 3 & 4) G L U C pg/ml (2) * 1 (exp 1) 2 (exp 3 & 4) LH ng/ml (1) 1 (exp 2)

NSB%

3.1

5.2 3.7 6.1

4.1 2.6 4.1

3.6 1.7 3.0 1.3

3.6 1.0 0.8

3.2 3.7

1.5

B0%

42.3

22.1 28.3 24.8

19.9 40.5 40.0

40.9 26.4 37.6 33.5

25.8 35.2 32.3

39.4 66.3

18.7

Detection limit

0.24

0.11 0.05 0.03

0.60 0.30 0.16

0.18 0.16 0.23 0.20

0.44 0.48 0.33

22.8 91.4

0.03

High pool

5.29±0.39

1.10+0.189 1.38±0.137 0.60+0.029

3.03±0.052 5.23±0.426 4.27±0.033

9.43±0.329 5.06±.535

-5.84±0.324

5.14±0.661 0.81±0.024 0.66±0.031

142+4.4 962±17.5

0.80±0.012

Medium pool

1.40±0.05

-0.65±0.054 0.40±0.040

2.18+0.211 3.45+0.058

-

--

3.40±0.132 -

-268±9.0

0.29±0.007

L o w pool

1.02±0.08

O.O6±O.0O3 0.44±0.081 0.16±0.030

0.36+0.045 2.76±0.159 1.10+0.173

1.98±0.184 0.24±0.023 0.89±0.300 1.14±0.071

1.21+0.661 0.65±0.028 0.44±0.036

39+2.4 130±7.8

0.053±0.003

PRL = prolactin, T3 = triiodothyronine, T4 = thyroxine, C O R T = corticosterone, INS = insulin, G L U C = glucagon, L H = luteinising hormone. The number of assays performed for each hormone is given in brackets. An * indicates that different pools were used for all the assays performed.

Table 2.3: The intra-assay coefficients of variation for the hormone assays performed.

Assay

PRL ng/ml (1) 1 (exp 3) T3 ng/ml (3) * 1 (exp 1) 2(exp 2) 3 (exp 3 & 4) T4 nM/L (3) * 1 (exp 1) 2 (exp 2) 3 (exp 3 & 4) C O R T ng//il (4) * 1 (exp 2A) 2 (exp 2B) 3 (exp 3 & 4) 4 (exp 3 & 4, repeats) INS ng/ml (3) * 1 (exp 1) 2 (exp 2) 3 (exp 3 & 4) G L U C pg/ml (2) * 1 (exp 1) 2 (exp 3 & 4) L H ng/ml (1) 1 (exp 2)

High Pool (%)

14.6

29.8 16.9 8.4

3.0 14.0 1.3

6.9 21.1 -

11.0

22.3 5.9 9.3

6.16 3.62

2.61

Medium Pool (%)

7.16

-14.3 17.0

16.6 2.9 -

--7.6 -

-6.69

4.03

Low Pool (%)

14.9

13.6 30.9 31.3

17.7 10 27.2

18.4 18.6 66.8 12.4

5.82 7.9 16.9

12.1 11.9

10.8

PRL = prolactin, T3 = triiodothyronine, T4 = thyroxine, C O R T = corticosterone, INS = insulin, G L U C = glucagon, LH = luteinising hormone. The number of assays performed for each hormone is given in brackets. An * indicates that different quality controls were used for all the assays performed.


10. Molecular techniques

Reagents

0.1% DEPC water

Luria Bertani (LB) media

LB selective media

LB selective plates

WxTAE

WxTBE

5 % acrylamide gel

1 ml of diethylpyrocarbonate (DEPC) in 1 L of nanopure

water, mixed thoroughly and incubated at room

temperature overnight. Autoclaved for 20 minutes at 15

lb/sq inch on liquid cycle to inactivate DEPC.

To 950 ml nanopure add 10 g tryptone, 5 g yeast extract

and 10 g NaCl, adjust volume to 1 L. Sterilise by

autoclaving 20 minutes at 15 lb/sq inch on liquid cycle.

Make LB as described. Following autoclaving cool with

agitation in a water bath to 60°C and add 100 ug/ml of

ampicillin.

To 950 ml nanopure add 10 g tryptone, 5 g yeast extract,

10 g NaCl and 15 g agar, adjust volume to 1 L. Sterilise

by autoclaving 20 minutes at 15 lb/sq inch on liquid cycle.

Cool with agitation in a water bath to 60°C and add 100

figlnA of ampicillin. Pour into sterile plastic petri dishes,

allow to set and expose to U V light overnight.

Refrigerate.

48.4 g tris base, 11.42 ml glacial acetic acid and 20 ml of

0.5 M E D T A p H 8. Adjust volume to 1 L with nanopure

water.

109 g tris base, 55 g boric acid and 40 ml of 0.5 M E D T A

p H 8. Adjust volume to 1 L with nanopure water and

sterilise by autoclaving.

14.4 g urea, 3 ml 10 x TBE, 3.8 ml 4 0 % acrylamide

(acryhbis-acryl = 19:1), add 8 ml of dH20, stir at room

temperature until dissolved, adjust volume to 30 ml and

degas. While stirring add 240 ul of 1 0 % ammonium

persulfate in d H 2 0 and 32 ul of T E M E D (Bio Rad). Pour

gel immediately.

R N A extraction

R N A was isolated from emu liver, adipose, mediobasal hypothalamus and preoptic area

using a method based on that developed by Chomczynski and Sacchi (1987), and a


commercial extraction buffer, RNAzol B (Tel-Test Inc.). Samples were ground in

liquid nitrogen using a mortar and pestle resting on dry ice, to prevent RNase activity.

The powder was then transferred to an appropriate volume of extraction buffer and

processed according to the directions supplied. The methodology was modified to

maximise the extraction of R N A from emu adipose, brain and liver tissues. The

complete methodology is given below.

RNAzol B RNA extraction protocol

Approximately 1 g of the sample tissue was ground to a powder in liquid nitrogen with

a mortar and pestle resting on dry ice (for brain tissues the volume of RNAzol was

reduced to 5ml). The powder was transferred to a glass grinder containing 10 ml of

RNAzol B and homogenised. The homogenate was transferred to a baked 15 ml Corex

tube and rested on ice for 5 minutes. A volume of 2 ml of chloroform was added, and

the tube vortexed to ensure thorough mixing, the tube was placed on ice for a further 5

minutes. Samples were centrifuged at 12,000 x g and 4°C for 1 hour. The upper

aqueous phase was pipetted into a sterile 15 ml glass Corex tube, with care taken not to

disturb the interphase, an equal volume of isopropanol was added and mixed by

vortexing. Samples were stored at -20°C for more than 45 minutes (for adipose and

brain tissues that do not have abundant mRNA's, samples were stored between 24 and

48 hours to maximise the precipitation of R N A ) . Tubes were centrifuged at 6,000 x g

and 4°C for 45 minutes. The supernatant was removed and the pellet rinsed once with 5

ml of 7 5 % ethanol, followed by centrifugation at 6,000 x g and 4°C for 45 minutes. The

supernatant was removed and the pellet air dried for 10-15 minutes. The pellet was

dissolved in 1 ml (liver), 0.1 ml (adipose) or 20 ul (mediobasal hypothalamus/preoptic

area) of D E P C treated distilled water, with the addition of 1 ul of RNasin RNase

inhibitor (Promega) to each sample, and stored at -70°C.

Ribonuclease Protection Assay (RPA)

RPA's for neuropeptide Y (NPY) and vasoactive intestinal peptide (VIP) expression in

the mediobasal hypothalamus and preoptic area were performed at the Roslin Institute,

Scotland, with the assistance of Dr Boswell (Chapters 5 and 6). The technique was then

established in our laboratory with the generous gift of emu specific N P Y , VIP and p-

actin clones. The R P A technique was subsequently used to determine the expression of

N P Y and VIP in the final experiment (Chapter 7).


Total R N A was extracted from tissues using the methodology given above, and stored

under 7 0 % ethanol before being sent to the Roslin Institute (Chapters 5 and 6) or used in

our laboratory (Chapter 7). At Roslin, emu specific VIP, N P Y and p-actin inserts were

ligated into the bluescript plasmid (Stratagene) and transformed into the XLl-blue

Escherichia coli strain (Stratagene), to be used as probes. The Ambion R P A III kit

was used to determine N P Y and VIP expression using the methodology below:

Probe and control preparation:

Cultures of clones containing VIP, N P Y and p-actin were grown up overnight in 5 or 10

ml of L B selective media. The vector D N A was extracted using Q I A G E N mini-preps

(Roslin) or midipreps ( U W A ) following the manufacturers directions.

Table 2.4: RNA polymerase and restriction enzyme required for each probe to obtain sense and antisense strands.

Probe Antisense strand Sense strand

E m u VIP T7; Eco RI T3; Hind HI

E m u N P Y T7; Eco RI T3; Hind III

E m u (3-actin T3; Hind III T7; Eco RI

Restriction Digestion:

The plasmid D N A obtained was digested by adding 20 ul buffer and 10 ul enzyme

(Boehringer Mannheim see Table 2.4) tol80 ul plasmid D N A (-40 ug). This was then

vortexed and microfuged briefly. The tubes were then incubated at 37°C for 60-90

minutes after which a further 5 ul of enzyme was added and the tubes incubated for a

further 2 hours. At this point the samples were either frozen at -20°C or phenol-

chloroform extraction performed.

Phenol-chloroform extraction:

To each 200 ul digest 100 ul phenol and 100 ul of the chloroform mix (24:1,

chloroform:isoamyl alcohol) was added. The tubes were vortexed for 1 minute and

microfuged for 2 minutes at maximum rpm. The upper aqueous layer was transferred to

a new tube. A volume of 200 ul of d H 2 0 was added to the original tube and the

contents were back extracted by vortexing for 1 minute and microfuging for 2 minutes

at maximum rpm. The upper aqueous layer was transferred to the tube containing the

first upper layer removed. To this was added 400 ul of the chloroform mix and the tubes


were vortexed for 1 minute and microfuged for 2 minutes at maximum rpm. The upper

aqueous layer was transferred to a new tube and the chloroform extraction repeated. To

precipitate the digested plasmid D N A 40 ul of 3 M sodium acetate p H 5.2 and 1100 ul

of cold 100% ethanol were added. The tubes were vortexed and incubated at -20°C for

at least 30 minutes. To pellet the D N A the tubes were microfuged for 20 minutes at 4°C

and maximum rpm, and rinsed with 7 0 % ethanol (1 ml added and tubes microfuged for

10 minutes at maximum rpm). The supernatant was removed completely and the pellet

air dried for approximately 5 minutes. The sense and antisense plasmid D N A templates

were then reconstituted in 32 ul of dH20 and the concentration determined

spectrophotometrically.

Preparation of positive control from the sense strand for each probe:

A total of 1 ug of sense template D N A was denatured for 3 minutes and placed on ice.

A nucleotide mix was prepared by combining 2 fi\ of each supplied nucleotide with 2 ul

of dtÔ. To each of the templates 1 fi\ of the nucleotide mix, 2 ul of buffer and 2 ul of

the appropriate R N A polymerase (Table 2.4) were added and the final volume was

adjusted to 20 ul with dHÔ. The reactions were incubated for 2 hours at 37°C and 2 ul

of DNase was added and the tubes incubated for a further 20 minutes at 37°C. To stop

the reaction and precipitate the sense strand 2 ul of E D T A (0.2 M ) , 2.5 ul of 4 M

lithium chloride (LiCl) and 75 ul of 100% ethanol were added to the tubes. These were

vortexed and incubated at -70°C for greater then 30 minutes or overnight at -20°C. The

tubes were microfuged for 20 minutes at 4°C and as much supernatant as possible

removed before rinsing with 7 0 % ethanol. The sense strands were reconstituted in 40 fi\

of deionized formamide and incubated at 70°C for 1 hour. To use as a control 1 ul of

sense strand was diluted 1:5000 with dtÔ immediately prior to performing the

reaction.

Labeling of probes:

To 0.5 fil of the antisense template D N A , obtained as described above, 0.5 fi\ nucleotide

mix (1 ul each of ATP, CTP, G T P and a 1:200 dilution of U T P + 6 ul of dH 20), 1 ul

buffer, a trace amount of polymerase (T7 N P Y and VIP; T3 p-actin) and 6.75 ul dlfcO

was added. The tubes were microfuged briefly and 1.25 ul [oc-32P]UTP (800 Ci/mmol)

was added. The tube contents were mixed by gentle pipetting and incubated for 30

minutes at 37°C. A volume of 1 ul of DNase was added and the tubes incubated for 20

minutes at 37°C. The reaction was stopped and the antisense R N A precipitated by


adding 1 ul of 0.2 M E D T A , 1.25 ul of 4 M LiCl and 37.5 ul of cold 1 0 0 % ethanol.

The tubes were then incubated at -20°C for more than 15 minutes. To pellet the R N A

the tubes were microfuged for 15 minutes at 4°C and maximum rpm and the supernatant

completely removed. The pellet was rinsed with 1 ml of 7 0 % ethanol and microfuged

for 10 minutes, the supernatant was then completely removed and the R N A

reconstituted in 100 ul of sterile dH 2 0. A volume of 2 ul of this solution was added to

scintillant and counted using a beta counter.

Hybridisation:

The hybridisation reagents listed in Table 2.5 were added to a fresh eppendorf tube. The

tube contents were mixed thoroughly and incubated at -20°C for more than 15 minutes.

The tubes were microfuged at maximum rpm for 15 minutes at 4°C and the supernatant

was completely removed. The pellet was resuspended in 10 ul of hybridisation buffer,

vortexed and briefly microfuged. Samples were denatured at 90-95°C for 3-4 minutes

and then incubated overnight at 42°C.

Table 2.5: Reagents for RPA hybridisation.

Reagent

Sample R N A (itg)

Yeast R N A (itl)

1:5000 dilution sense

strand (pi)

d H 2 0 (pi)

N P Y Probe

VIP probe

[3-actin probe

5 M N H 4 0 A c (itl)

100% ethanol (pi)

Blank

-

2

-

28

>50,000 cpm

>50,000 cpm

>50,000 cpm

1/10 volume

2.5 volumes

+ve control

-

2

1

27

>50,000 cpm of

the appropriate

probe

1/10 volume

2.5 volumes

sample

10-20

-

-

to 30 itl

>50,000 cpm

>50,000 cpm

>50,000 cpm

1/10 volume

2.5 volumes

RNase digestion of hybridized probe and sample RNA:

The RNaseA/RNaseTl was diluted 1:100 with RNase digestion buffer, vortexed and

briefly microfuged to ensure even dispersal of components. The hybridisation reactions

were briefly microfuged to collect all liquid at the bottom of the tube. A volume of 150

ul of the 1:100 RNaseA/RNaseTl dilution was added, vortexed and the tubes

microfuged briefly. The reactions were incubated for 30 minutes at 37°C. A volume of

225 fi\ of RNase inactivation/precipitation III solution was added, the tubes vortexed


and microfuged briefly. The reactions were incubated at -20°C for more than 15

minutes. Following incubation the reactions were microfuged at maximum rpm for 15

minutes at 4°C and the supernatant carefully removed. The pellets were resuspended in

8 ul of gel loading buffer n, vortexed vigorously and briefly microfuged. Reactions

were incubated at 90-95°C for 3 minutes, briefly vortexed and microfuged and rested on

ice. The samples were loaded onto a 1 x T B E , 5 % acrylamide gel (0.75 m m thick, 12

cm long x 15 c m wide) and run at a constant 250 volts for approximately 1 hour or until

the leading dye band neared the bottom of the gel. Once finished running the gel was

transferred to filter paper, the origins and orientation marked, covered with plastic wrap,

and dried. Once dry the gel was exposed to a phosphoimaging plate overnight at -80°C.

The bands corresponding to B-actin, N P Y and VIP were visualised and analysis of their

intensity was performed on a Macintosh (iMac) computer using the public domain NIH

Image program version 1.61 (developed at the U.S. National Institutes of Health, an

available on the Internet at http://rsb.info.nih.gov/nih-image/).

http://rsb.info.nih.gov/nih-image/

58

Chapter 3

The emu leptin gene.

1. Introduction

2. Materials and Methods

3. Results

4. Discussion

Page

59

61

75

77

Chapter 3 - The emu leptin gene 59

1. Introduction

In emus, reproduction is associated with decreased appetite and adiposity. This could be

expected to result in parallel decreases in the circulating concentration of leptin. In

humans and rodents decreases in the concentration of leptin would stimulate appetite. This

clearly does not occur in the emu, since anorexia and diminished appetite are features of

incubating and non-incubating animals respectively during the breeding season (Mawson,

1992; Blake, 1996; Blache & Martin, 1999; Blache et al, 2001b). So h o w is appetite

suppressed in emus despite diminishing adipose reserves and what role, if any, does leptin

play?

In species that initiate reproduction in response to decreasing day length, seasonal changes

in body weight and adiposity, are paralleled by changes in the concentration of leptin

(Bocquier et al, 1998; Atcha et al, 2000; Horton et al, 2000; Mercer et al, 2000a;

Concannon et al, 2001; Marie et al, 2001). It has been proposed that higher sensitivity to

leptin during short days increases the responsiveness to food deprivation and enhance the

ability to mobilise adipose reserves in sheep (Bocquier et al, 1998). In the Siberian

hamster, leptin administration decreased fat reserves in animals exposed to short but not

long photoperiods and had no effect on food intake under either photoperiod (Atcha et al,

2000). The decrease in fat reserves during short photoperiod indicates enhanced sensitivity

to the actions of leptin. In this species, food deprivation during short but not long

photoperiods decreased the leptin concentration and leptin receptor (Ob-Rb) expression

indicating a greater sensitivity to food deprivation (Mercer et al, 2000a).

The involvement of leptin in appetite and adiposity control during the reproductive period

could take several forms. From the information available on leptin in other species and our

knowledge of the appetite and adiposity changes that occur there are four scenarios that

seem likely. First, as adiposity decreases, elevated leptin concentrations are maintained

and act to depress appetite. Second, that the leptin concentrations may decrease in parallel

with adiposity, and this fails to stimulate appetite as a result of the actions of other factors,

possibly related to photoperiod, influencing appetite. Third, that changes in the expression

of the leptin receptor at the hypothalamic level alter the responsiveness to leptin. Finally,

it is possible that emus do not have the leptin gene or its product. However, this final

scenario is unlikely based on evidence for the existence of the leptin gene in chickens.

First, southern hybridisations of the mouse leptin gene to chicken D N A indicated


homology between the two species (Zhang et al, 1994). Second, sequences for the

chicken leptin gene and chicken leptin receptor have been published (Taouis et al, 1998;

Ashwell et al, 1999a; Horev et al, 2000). As leptin is highly conserved in mammalian

species it is likely that it is also highly conserved in avian species. As such, the existence

of the leptin gene in chickens argues strongly for the existence of a homologous sequence

in the emu.

A bioassay for chicken leptin was recently developed by Dr Friedman-Einat based on a

human leptin bioassay produced by the Merck Company (Rosenblum et al, 1998). It

utilises a firefly luciferase reporter construct and the chicken leptin receptor transfected

into the HEK-293 embryonic kidney cell line (Rosenblum et al, 1998). Activation of the

leptin receptor by leptin in plasma or serum in turn activates STAT3, a transcription factor,

causing the reporter construct to emit light. The amount of light emitted can be quantified

and the activity of leptin in the sample deduced. It could be expected that if emu and

chicken leptin sequences have been highly conserved that leptin in emu serum will activate

the chicken leptin receptor in this assay.

To demonstrate the existence of leptin in emus and in particular its presence in serum,

which is the first step in the study of the possible role of leptin in the emu, w e tested the

following hypotheses;

1. Emus have a leptin gene and its sequence is homologous to the published chicken and

mammalian sequences, and

2. E m u serum will activate the chicken leptin receptor in the chicken leptin bioassay.



Experimental design

The samples used for this experiment were obtained from male and female emus that had

been used in observational studies. Several molecular techniques were employed to test

the hypotheses outlined in the introduction. First, R N A was extracted from liver and

adipose tissue and used or reverse transcription polymerase chain reaction (RT-PCR) with

the primers for the chicken leptin gene sequence. Rapid amplification of c D N A ends

(RACE) was performed to obtain the complete emu leptin gene sequence. Second, chicken

and mammalian leptin probes, were used to screen an emu cosmid library for homologous

sequences. Finally, emu serum was added to a cell line containing the chicken leptin

receptor to determine if components within the serum could activate the chicken leptin

receptor.

Animals and samples

The tissue samples used for this work were obtained from emus of different sexes, at

various times of the year. All of the animals were in good health when they were

euthanased. The serum samples used in the chicken leptin bioassay were obtained from six

male emus used in observational studies by another researcher. These samples were

obtained over the course of a year and all of the animals were in good health.

Poly A+ RNA extraction

Poly A + R N A was obtained using the protocol described by Sambrook et al (1989).

Reverse Transcription Polymerase Chain Reaction (RT-PCR)

The cloning of the chicken leptin gene was published after the commencement of m y Ph.D.

(Taouis et al, 1998; Ashwell et al, 1999a). Primers derived for chicken leptin and a

chicken leptin probe (supplied by Dr Ashwell) were subsequently used in attempts to clone

the emu leptin gene (Ashwell et al, 1999a).

Reverse Transcription (RT)

c D N A was obtained from R N A using M - M u L V reverse transcriptase and 10 x reverse

transcription (RT) buffer (New England BioLabs) following the method supplied by the

manufacturer. For the initial isolation of a fragment of the leptin gene, a specific reverse

primer designed by Nakavisut (1998) (OBR1 - 5 V - T A C T C C A C A G A G G T G G T G G C - 3 V )


was used in the R T reaction. This primer was designed from a region of high homology

between all published D N A sequences for the mammalian leptin gene, and also high

homology to the published chicken sequence. The reagents listed in Table 3.1 were added

to a fresh 200 ul eppendorf tube and incubated at 37°C for 1 hour. The reaction was

heated to 90°C for 5 minutes and then placed on ice. These steps were performed using a

Hybaid OmniGene thermal cycler. The dNTP's were obtained from Pharmacia Biotech

and the RNasin from Promega.

Table 3.1: Volumes of reagents used in the reverse transcription reactions.

Reagent Volume (pi)

10 x RT buffer 5.0

R N A volume to give 2 p%

dNTP's (25 m M ) 2.0

OBRl(100pmol//tl) 0.5

RNasin (40 units//d) 1.0

M - M u L V (25 units/jtl) 4.0

dH20 to a final volume of 50 pi

Polymerase Chain Reaction (PCR)

The P C R was performed using the external sense primer sCLepl (5V-

C A C C A G G A T C A A T G A C A T T T C A C - 3 V ) (Ashwell et al, 1999a) and the O B R 1 primer

(Nakavisut, 1998). The reagents listed in Table 3.2 were added to a fresh 200 ul eppendorf

tube, adding the Taq D N A polymerase (Promega) last. The 10 x PCR buffer (MgCl2 free)

was supplied with the Taq D N A polymerase.


Table 3.2: Volumes of reagents used in the polymerase chain reactions.

Reagent Volume (pi)

R T product 10.0

OBR1 (100 pmoVpl) 0.1

sCLepl(10pmol//d) 0.5

10 x PCR buffer 5.0

Taq D N A polymerase (5 units/jtl) 0.5

dH 20 33.9

Total 50.0

Thermal cycling was performed using a Hybaid OmniGene thermal cycler under the

conditions given in Table 3.3.

Table 3.3: Thermal cycling conditions used for the polymerase chain reaction.

Step Denature Anneal Extend Number of cycles

1 94°C, 1 min 50°C, 1 min 72°C, 2 mins 5

2 94°C, 30 sec 50°C, 30 sec 72°C, 1 min 30

3 . - 72°C,5mins 1

The products were cooled to 15°C in the thermal cycler before being run on a 1% agarose

gel or used directly in nested PCR.

Nested PCR was performed using the primers:

sCLep2 (5V-CGTCGGTATCCGCCAAGCAGAGGG-3V) and

aCLep2 (5V-CCAGGACGCCATCCAGGCTCTCTGGC-3V) (Ashwell et al, 1999a).

The volumes of the nested PCR reagents used, dNTP's (Pharmacia Biotech), 10 x PCR

buffer, MgCl2, Taq D N A polymerase (Promega), and Triton x 100 (Sigma), are given in

Table 3.4.


Table 3.4: Volumes of reagents used in nested PCR.

Reagent Volume (pi)

PCR product

dNTP's (25 m M )

sCLep2(10pmol//d)

aCLep2(10pmol//tl)

10 x PCR buffer

MgCl2 (25 m M )

Taq D N A polymerase (5 units//tl)

Triton x 100 (diluted 1:100)

dH20

Total

1.0

0.1

0.1

0.1

1.0

0.7

0.1

0.5

6.4

10.0

Thermal cycling was performed as described in Table 3.3 with an annealing temperature of

55°C in steps 1 and 2.

Purification of PCR products

O n the basis of the published leptin gene sequences a 260 base pair (bp) P C R product was

expected using these primers. The P C R products were visualised using 1 % agarose gels,

excised from the gel using a sterile razor blade and extracted with the Qiaex II gel

extraction kit (QIAGEN) according to the manufacturers directions.

Cloning

Reagents

Luria-Bertani (LB) media and plates (with or without ampicillin)

The media and plates were made according to the directions of Sambrook et al

(1989).

Competent cells

Competent cells were prepared according to the methodology of Sambrook et al (1989).

Briefly, Escherichia coli (E. coli) of the X L 1-blue strain were streaked onto L B media

plates (no ampicillin) and incubated overnight at 37°C. A single colony was selected from

the plate and used to inoculate 5 ml of L B medium. The culture was grown overnight at

37°C with gentle agitation. This culture was diluted 1:100 with 500 ml of L B media pre-


warmed to 37°C and was incubated at 37°C with vigorous shaking to an OD650 of 0.8. The

cells were chilled quickly by swirling gently in an ice water bath for approximately 5

minutes. The cells were centrifuged at 8000 rpm for 8 minutes at 4°C. The supernatant

was poured off and the pellet resuspended in 125 ml of ice-cold 0.1M M g C l 2 and

immediately centrifuged as described above. After pouring off the supernatant the cells

were resuspended in 125 ml of ice-cold 0.1M CaCl2 and incubated on ice for 20 minutes,

before being centrifuged as described above. After pouring off the supernatant the cells

were resuspended gently in 25 ml of ice-cold 0.1M CaCl2 containing 1 4 % v/v glycerol, and

the suspension was incubated on ice for less than 30 minutes. The competent cells were

aliquoted in 0.2 ml volumes into pre-chilled eppendorf tubes and snap frozen in liquid

nitrogen. The cells were then stored at -80°C.

Ligation of the purified PCR product into the pGEM-T easy vector.

The purified nested P C R product was run on a 1 % agarose gel and a visual assessment

with Kodak ID20 software was used to estimate the concentration. Ligation of the purified

product into a vector was performed using the p G E M - T easy kit (Promega) (Table 3.5).

A n insert to vector ratio of 3:1 was used where possible.

Table 3.5: Volumes of reagents used in the ligation reaction.

Reagent Volume (pi)

10 x ligase buffer 1

Ligase (3 units/pl) 1

pGEM-T (50 ng/pl) 1

insert -150 ng

(1H20 to a final volume of 10 pi

The ligation was prepared on ice and allowed to proceed either overnight at 4°C, or at

15°C for a minimum of 2 hours.

The ligated product was transformed into the E. coli strain XL-1 blue using the heat shock

method (Sambrook et al, 1989). Briefly, competent cells were thawed on ice and 2 ul of

the ligation reaction was added. The reaction was incubated on ice for 30 minutes. The

reaction tube was transferred to a waterbath at 42°C for 90 seconds and returned to ice for


1-2 minutes. A volume of 800 ul of L B media was added to the reaction tube and

incubated at 37°C for 90 minutes in a shaking incubator. The reaction was microfuged at

5000 rpm for 5 minutes and 800 ul of L B media was removed with a pipette. The pellet

was resuspended in the remaining L B media and diluted to 1:10 and 9:10, and spread onto

L B plates containing ampicillin. The dilutions used allowed individual colonies to be

selected after the incubation period. The presence of ampicillin in the media allowed E.

coli colonies containing the p G E M - T vector to be identified as it confers ampicillin

resistance. The inoculated plates were incubated at 37°C overnight.

Colony PCR.

Colony P C R was used to determine colonies that harboured inserts of the desired size.

Briefly, individual colonies that had grown on the selective plates were touched with a

sterile sealed glass pasteur pipette (Chase Instruments), which was then touched to a fresh

selective plate with an identifying grid drawn on the petri dish. After this the tip of the

glass rod was placed in an eppendorf tube with 10 ul of sterile distilled water and agitated

gently to produce a bacterial suspension. The tube was numbered to correspond with the

colony's grid location on the new plate. The SP6 and T7 primers were used to identify

colonies with inserts of the correct size, as they anneal either side of the multiple cloning

region of the p G E M - T vector. The volumes of P C R reagents, dNTP's (Pharmacia

Biotech), 10 x P C R buffer, M g C l 2 and Taq D N A polymerase (Promega), used in the

colony P C R are given in Table 3.6.

Table 3.6: Volumes of reagents used for colony PCR.

Reagent Volume (pi)

Bacterial suspension 1.0

dNTP's (25 m M ) 0.1

SP6(10pmol//d) 0.1

T7(10pmol/ftl) 0.1

10 x PCR buffer 10

MgCl2 (25 m M ) 0.7

Taq D N A polymerase (5 units/pl) 0.1

dH20 6.9

Total 10.0


The thermal cycling conditions for the colony P C R are as given in Table 3.3, with the

exception that the annealing temperature used at step 2 was 55°C. The colony P C R

products were run on a 1 % agarose gel and those ascertained to have inserts of the desired

size were grown up in 5 ml of LB + ampicillin media (mini culture) for about 8 hours.

This was diluted 1:100 into 25 ml of LB + ampicillin media (maxi culture) and grown up

overnight. The remainder of the mini culture was used to make glycerol stocks by adding

500 ul of the culture and 500 ul of 8 0 % glycerol (Sigma) to a sterile eppendorf tube that

was then stored at -80°C. The plasmid midifilter kit (QIAGEN) was used to obtain

plasmid D N A according to the manufacturer's directions from the maxi culture. After

assessing the resulting plasmid D N A concentration and quality spectrophotometrically, it

was sequenced at the Immunology Department of Royal Perth Hospital. The GeneJockey

II program was utilised to find regions of the sequence corresponding to SP6, T7 and the

primers used in the nested PCR. Once the regions amplified by these were found they

were compared to other known leptin gene sequences with the GeneJockey II program and

also utilising the B L A S T service offered by the National Institute of Health, USA,

accessed via the internet (http://www.ncbi.nlm.nih.gov/).

Cosmid library screening

A n emu cosmid library (Stratagene) (Hammond, 2000) was screened with a 260 bp

mammalian leptin fragment and Ashwell's chicken leptin fragment labelled with [a" P]

dCTP(NEN).

Restriction digestion of plasmid DNA

The plasmid D N A of both leptin fragments were digested with restriction enzymes to

remove the vector D N A as this shares homology with cosmid D N A used in the emu c D N A

library. The volumes of EcoRl, 1 x N E buffer for EcoRl (New England Biolabs), BSA,

Ncol, Sail and 10 x buffer D (Promega) used in the restriction digest are given in Table

3.7. Ashwell's chicken leptin fragment was digested with £coRI and the mammalian

fragment with Ncol and Sail by adding the reagents as indicated in Table 3.7, mixing them

well by pipetting and incubating overnight at 37°C.

http://www.ncbi.nlm.nih.gov/


Table 3.7: Volumes of reagents used for restriction digests of plasmids containing chicken or mammalian leptin fragments.

Reagent Chicken leptin (pi) Mammalian leptin (pi)

Plasmid D N A (10 pg) 17 16

BSA (acetylated 10 mg/ml) - 0.5

EcoRl 1.5

Ncol - 3

Sail - 3

1 x N E buffer for EcoBl 5

10 x buffer D - 5

Distilled water 26.5 12.5

Total 50 50

The restriction digest products were precipitated by adding 5 ul of 3 M sodium acetate (pH

5.2) and 165 ul of absolute ethanol. The reagents were mixed thoroughly and stored at -

20°C for at least one hour. The reactions were centrifuged at 12,000 g and the supernatant

removed completely. The pellets obtained were resuspended in 15 ul of sterile distilled

water. The entire volume was then loaded onto a 1 % agarose, 1 x T B E gel and run for

approximately one hour at 80 volts. The 100 bp marker (Promega) was also run with the

restriction products and a band of approximately 300 bp was excised and purified with the

Qiaex II gel extraction kit (QIAGEN) as described previously.

Labelling probes

The chicken and mammalian leptin probes were labelled using the N e w M E G A P R I M E kit,

protocol 1606 (Amersham). Briefly, the insert D N A concentration was diluted to 5 ng/ul

with distilled water. Using this dilution 25 ng of the D N A insert was added to a sterile

eppendorf tube with 5 ul of the supplied primers, and denatured by heating to 95-100°C for

5 minutes in a water bath. The tube was microfuged briefly. At room temperature, 10 ul

of labelling buffer, 2 ul of enzyme and 23 ul of distilled water were added to the tube. The

tube was microfuged briefly and 5 ul of [a"32P] d C T P (NEN), specific activity 3000

Ci/mmol, was added and mixed by gentle pipetting, followed by brief microcentrifugation.

The tube was incubated at 37°C for 10 minutes. The reaction was stopped by the addition

of 5 ul of 0.2M E D T A and denatured by heating to 95-100°C for 5 minutes. The labelled

insert was chilled on ice before being put through a N I C K column (Pharmacia Biotech) to


remove unincorporated nucleotide, according to the manufacturers directions.

Approximately 100 ul fractions were collected and those with the highest levels of

radioactivity were identified with a geiger counter (Morgan series 900 mini monitor) and

pooled.

Southern Hybridisation

Reagents

Salmon sperm (SS) DNA

The SS D N A was prepared according to the directions of Sambrook et al (1989).

10 x PIPES (piperazine-N,N'-bis[2-ethane sulfonic acid])

To 700 ml of sterile distilled water 30.24 g of PIPES was added, and the pH was

adjusted to 6.5 with 0.1 N HC1 if required. To this was added 233.76 g of NaCl.

Sterile distilled water was added to give a final volume of 1 L and a final

concentration of 4 M NaCl and 0.1 M PIPES (pH 6.5).

20xSSC

To 800 ml of distilled water 175.3 g of NaCl and 88.2 g of sodium citrate were

added. The pH was adjusted to 7 with 10 N N a O H and distilled water added to

give a final volume of 1 L with. The solution was sterilised by autoclaving on a

liquid cycle (15 lb/sq inch) for 20 minutes.

10% SDS

To 900 ml of distilled water 100 g of electrophoresis grade SDS was added. The

solution was heated to 68°C to assist dissolution. The pH was adjusted to 7.2 with

concentrated HC1 and distilled water added to give a final volume of 1 L.

1M Tris-Cl (pH 7.6)

To 800 ml of distilled water 121.1 g of Tris base was added, the pH was adjusted to

7.6 with concentrated HC1 and distilled water added to give a final volume of 1 L.

The solution was sterilised by autoclaving on a liquid cycle (15 lb/sq inch) for 20

minutes.


1M Tris-Cl (pH 7.6) + 1.5MNaCl

Made as for 1 M Tris-Cl (pH 7.6) with the addition of NaCl to a final concentration

of 1.5 M.

O.SMNaOH

To 800 ml of distilled water 20 g of N a O H was added. This was allowed to

dissolve and the final volume was adjusted to 1 L with sterile distilled water.

Cosmid prehybridisation solution (2 x PIPES, 50% deionizedformamide, 0.5%

SDS(wA))

A volume of 200 ml of 10 x PIPES, was added with 500 ml of deionized

formamide and 50 ml of 10% SDS to a sterile 1 L Schott bottle. The final volume

was adjusted the to 1 L with sterile distilled water.

2xSSC + 0.1%SDS

This solution was prepared by diluting 20 x SSC 1 in 10, and 10% SDS 1 in 100

with sterile distilled water to give a final volume of 1 L in a sterile Schott bottle.

Ix SSC + 0.1% SDS

This solution was prepared by diluting 20 x SSC 1 in 20, and 10% SDS 1 in 100

with sterile distilled water to give a final volume of 1 L in a sterile Schott bottle.

Southern hybridisation to the emu cosmid library

The emu cosmid library was spread on to LB 4- ampicillin media plates in 200 fi\ volumes

at 1 x 106 and 1 x 107 dilutions, and grown up overnight at 37°C. Replica membranes were

made by placing a Hybond-N+ (Amersham) membrane on each plate and allowing them to

wet through. The membranes were then removed and placed colony side up onto fresh LB

+ ampicillin media plates and grown for a further four hours at 37°C.

Positive control membranes were prepared by denaturing plasmid DNA containing the

mammalian and chicken leptin gene sequences at 95°C for 5 minutes before dotting onto

Hybond-N+ membrane (Amersham). The membrane was allowed to air dry and then

baked at 80°C for one hour to fix the DNA.


The replica membranes were prepared for hybridisation by placing them colony side up for

30 seconds onto a series of number 3 Whatman papers (Whatman International Ltd)

prewetted with the following solutions:

1. 0.5 M NaOH,

2. 1 M Tris-HCl (pH 7.6), and

3. 1 M Tris-HCl (pH 7.6)+1.5 M NaCl.

The membranes were placed in a solution of 1 M Tris-HCl (pH 7.6) + 1.5 M NaCl and any

colony debris was gently removed by rubbing a gloved hand over the surface of the

membrane. The membrane was rinsed in fresh 1 M Tris-HCl (pH 7.6) + 1.5 M NaCl and

allowed to air dry on paper towel before being baked at 80°C for one hour.

The replica and positive control membranes were placed on fine nylon gauze prewetted

with the cosmid prehybridisation solution, this was gently rolled up and placed in Hybaid

H B - O V - B M hybridisation bottles. A volume of 20 ml of the cosmid prehybridisation

solution pre-warmed to 50°C was added to the bottle with 200 ul of salmon sperm D N A

that had been boiled for 10 minutes immediately prior to its addition. The hybridisation

bottle was sealed and the membranes were prehybridised at 46°C for a minimum of two

hours in a Hybaid mini oven. The probes were prepared as described previously and at the

completion of prehybridisation boiled at 95-100°C for 5 minutes and added to the

prehybridisation solution. Hybridisation was carried out overnight at 46°C in a Hybaid

mini oven. At the completion of hybridisation the solution was poured off and 20 ml of 2

x SSC + 0.1% SDS, pre-warmed to 60°C, was added to the bottles. The membranes were

washed for 5 minutes in a Hybaid mini oven, and then the solution was poured off and the

wash repeated twice. A final wash was performed for 30 minutes with 20 ml of 1 x SSC +

0.1% SDS pre-warmed to 60°C. A geiger counter (Morgan series 900 mini monitor) was

used to measure the background radiation on the membrane and the wash steps repeated if

this was deemed too high. The membranes were blotted on paper towel and wrapped in

cling film before being exposed to x-ray film (Kodak scientific imaging film) overnight at -

80°C. The film was developed using Kodak chemicals according to the manufacturers

directions.

Rapid amplification ofcDNA ends (RACE)

Following the amplification of an appropriately sized band from emu liver R N A with

leptin gene primers that were specific for the chicken (Ashwell et al, 1999a) Clontech's


Marathon R A C E c D N A amplification kit was employed to obtain the remainder of the 3'

and 5' ends of the sequence from poly A + emu liver R N A .

The protocols outlined for the Marathon kit were followed, with the exception of the

thermal cycling conditions, which were optimised for the Hybaid OmniGene thermal

cycler and are given in Table 3.8.

Table 3.8: The thermal cycling conditions for the Marathon RACE reactions.

Step Denature Anneal Extend Number of cycles

1 94°C, 30 sec - - 1

2 94°C, 30 sec 65°C, 1 min 68°C, 3 mins 5

3 94°C, 30 sec 60°C, 1 min 68°C, 3 mins 5

4 94°C, 30 sec 57°C, 1 min 68°C, 3 mins 30

5 - - 72°C, 5 mins 1

The four largest bands resulting from the R A C E reaction were excised from the gel and

purified using the Qiaex II kit (QIAGEN). P C R was performed on the products using the

nested primers reported by Ashwell et al (1999a) and the thermal cycling conditions

described in Table 3.3, with the exception that annealing at steps 1 and 2 were performed

at 55°C. Any fragments that were approximately 260 bp in length were ligated into

p G E M - T and transformed into the E. coli strain X L 1-blue, using the methodology

described previously. The resulting colonies were screened using the SP6 and T7 primers

to determine which clones contained inserts of the same size as the R A C E product. These

were screened further via the nested P C R reaction as above. Those colonies that contained

plasmids with inserts of the correct size were grown up to make glycerol stocks as

described previously and for plasmid D N A preparations using the midifilter kit (QIAGEN)

according to the manufacturer's directions. The quality and concentration of the plasmid

D N A were determined spectrophotometrically. Plasmids were sequenced at the

Immunology Department of Royal Perth Hospital.

Chicken leptin bioassay

This work was performed with Dr Friedman-Einat at the Institute of Agriculture, Volcani

Center, Israel. The bioassay utilises a firefly luciferase reporter construct and the chicken

leptin receptor transfected into the HEK-293 embryonic kidney cell line using methods


reported by Rosenblum et al (1998). The colonies transfected with the leptin receptor were

screened by Dr Friedman-Einat to detect the clone with the highest level of leptin-induced

luciferase activity. This clone (clepr210) was used to determine the biological activity of

leptin in serum samples from emus and chickens. The methodology is given briefly below:

Reagents

DMEM medium

1 0 % fetal calf serum, 50 ug/ml streptomycin and 50 U/ml penicillin

Growth medium

D M E M + 2 ug/ml puromycin

Culture plate preparation

Growth medium was gently aspirated from a culture stock of clepr210, leaving a layer of

cells. An aliquot of 300 fi\ of trypsin was added and allowed to spread across the plate.

Clumps of cells that formed were detached by gentle tapping. The cells were removed

from the bottom of the plate by adding 9 ml of D M E M medium and gently pipetting the

solution. An aliquot of 1 ml of the resulting solution was added to a new petri dish, and an

additional 8 ml of growth medium was added, mixed gently and incubated at 37°C in a

9 5 % air, 5 % carbon dioxide atmosphere. A volume of 18 ml of D M E M medium was

added to the remaining 8 ml of cell culture. While gently mixing this solution, aliquots of

500 ul (24 well Sarstedt) or 900 ul (6 well culture plate) were distributed into each well of

the culture plates (-50,000 cells/well). The plates were then incubated at 37°C in a 9 5 %

air, 5 % carbon dioxide atmosphere overnight.

Assessment of leptin activity in serum or plasma

The medium was aspirated gently and replaced with Optimem serum free medium

(GIBCO) pre-heated to 37°C (300 ul for 24 well or 600 ul for 6 well culture plates). The

serum samples (20-30 ul emu serum for 24 well or 100 ul for 6 well culture plates) were

tested in duplicate. A blank reaction was prepared in duplicate by adding only Optimem

serum free medium (GIBCO). The culture plates were incubated for 8-16 hours at 37°C in

a 9 5 % air, 5 % carbon dioxide atmosphere. The medium was poured off and allowed to

drain. Once dry, the cells were lysed by the addition of 100 ul of lysis buffer (Promega)

and vortexing (24 well culture plates). For the six well culture plates the same volume of


lysis buffer was used and after vortexing the bottom of the wells were scraped to ensure

that all cells were lysed. The lysed product was transferred to an eppendorf tube and

stored at -80°C. A n aliquot of 40 ul was removed for luminometric determination. Protein

determination was performed in triplicate with lul of the lysed product.

Luminometric determination

A volume of 40 ul of luciferase was added to the 40 ul aliquot of lysed cells immediately

prior to reading each sample. Tubes were tapped gently and quickly to mix and remove air

bubbles. The samples were then aliquoted into a sample tube and the light emitted was

quantified with a TD20e luminometer (Turner Design Inc.).

Protein determination

The protein concentration of each well was determined by Bradford assays using a 150-

fold dilution of each cell lysate. The assays were performed in duplicate. A standard

curve was set up using 0, 1, 2, 3, 4 and 5 ul of B S A (1 mg/ml) in duplicate.

Leptin receptor (LepR) induction

The luciferase activity and protein content measurements were used to calculate the

luciferase activity per fig of protein in each sample. These values indicate how much the

leptin receptors in the cell line were activated by leptin in the samples. To make

comparisons between individual bioassays the LepR induction of the samples were

expressed as proportions of their blank (Optimem serum free medium only) reaction

values. The calculations are as follows:

Sample LepR induction = (luciferase activity / ug of protein) / blank LepR induction

Blank LepR induction = luciferase activity / ug of protein

Data and statistical analysis

The significance of differences over the course of a year, were determined using the

Friedman two-way analysis of variance by ranks described by Siegel (1956). Non-

parametric statistical analysis was judged the most appropriate, as related samples were

used and the sample size was small (n = 6). The results are indicated as the mean ± s.e.m.


3. Results

RT-PCR

A leptin-like product was amplified from emu R N A . The homology of the emu derived

product was highly homologous with the ovine leptin gene sequence (98%).

Cosmid library screening

The chicken leptin fragment provided by Dr Ashwell failed to hybridise to the emu c D N A

cosmid library. A mammalian leptin gene fragment was used as a positive control in the

southern hybridisations. This showed a high degree of homology with Ashwell's chicken

fragment, even after high stringency washing.

RACE

Using either total or poly A + R N A from emu liver the sCLepl and aCLepl primers failed

to amplify any D N A fragments that resembled the leptin gene (Ashwell et al, 1999a). The

highly conserved mammalian O B R 1 primer and the chicken specific sCLepl primer also

failed to amplify the emu leptin gene sequence (Nakavusit, 1998; Ashwell et al, 1999a).

Chicken leptin bioassay

The emu serum samples used in the chicken leptin bioassay produced quantifiable amounts

of light, which indicates that components in the samples activated the chicken leptin

receptor (Table 3.9 and 3.10). Lower values for the leptin receptor induction were

obtained with emu samples than are obtained for the chicken. Samples collected from

emus during the course of a year indicated that the biological activity of leptin did not

differ in the emu over this period (Table 3.10).

Table 3.9: Leptin receptor induction of the chicken leptin bioassay by emu and chicken serum samples.

— ^ m u Chicken

Ratio of sample to blank 2.72 ±0.64 14.18 ±0.66 LepR induction Values are means ± s.e.m.

Chapter 3 - The e m u leptin gene

Table 3.10: Leptin receptor induction of the chicken leptin receptor bioassay by emu serum samples collected over the course of a year.

Summer Autumn Winter Spring Summer

Ratio of sample to blank 2.59 ±0.78 1.93 ±0.70 2.31 ±0.94 2.14 ±0.79 2.86 ±1.09 LepR induction

Values are means ± s.e.m.


4. Discussion

The first hypothesis, that emus have a leptin gene sequence, is supported by the results.

However, the results do not support a high degree of homology between the published

chicken and mammalian sequences. The second hypothesis is supported by the activation

of the chicken leptin receptor by components in emu serum. In addition activation of the

chicken receptor by emu serum provides the evidence that the leptin gene is present in the

emu. The ability of emu serum to activate the chicken leptin receptor also suggests that

emu and chicken leptins share a high degree of homology. A leptin-like product was

amplified using RT-PCR from both chicken and emu R N A that shared high homology with

the ovine leptin gene sequence (98%). However, ovine R N A and D N A are regularly used

in our laboratory, and other approaches such as R A C E and screening of the emu cosmid

library failed to identify emu leptin. As such, the product amplified by RT-PCR was

deemed to be the result of contamination from aerosols within the laboratory. The failure

of RT-PCR, R A C E and screening of the emu cosmid library to identify the emu leptin gene

would indicate that the emu sequence does not share a high degree of homology with

Ashwell et al's (1999a) chicken or the published mammalian leptin gene sequences.

Controls used while screening the emu cosmid library revealed a high degree of homology

between the mammalian sequence and Ashwell et al's (1999a) chicken leptin gene

sequence. This is unexpected, as generally genes are more highly conserved between

avian species than between avian and mammalian species. These results could be taken to

indicate that the published chicken leptin gene sequence is not correct. Other researchers

have also concluded that the published sequence for the chicken leptin gene is not correct,

and is instead a contamination artefact (Friedman-Einat et al, 1999; Varma, 2000; Dunn et

al, 2001). The problem of contamination artefacts being produced using the P C R

technique is well documented (Sarkar & Sommer, 1990; Erlich et al, 1991; Neumaier et al,

1998; Burkardt, 2000; Urban et al, 2000).

There are several approaches that can be taken to determine if the published chicken leptin

gene sequence is correct. First, sequencing of the chicken and emu leptin gene sequences

using alternative methods to those employed by Ashwell et al (1999a) and Taouis et al

(1998). A group led by Dr Friedman-Einat at the Volcani Centre is currently attempting

this using their chicken leptin receptor sequence to capture the protein ligand and obtain its

N-terminal amino acid sequence (Horev et al, 2000). Once the N-terminal amino acid


sequence is known degenerate oligo primers can be designed. PCRs can then be

performed using these primers to obtain the complete chicken leptin gene sequence.

Second, testing of the protein product of the published chicken leptin gene sequence in the

chicken leptin bioassay. The failure of this product to activate the receptor could indicate

that the published chicken leptin gene sequence is incorrect. Third, the sequence of the

emu leptin receptor gene could be determined. If a high degree of homology between the

leptin receptor genes from e m u and chicken is found, it will argue strongly for sequence

conservation of the leptin gene.

The activation of the chicken leptin receptor by components within emu serum was much

lower than observed for the chicken. This could be due to lower specificity of the assay for

emu leptin or to lower biological activity of emu leptin. The bioassay also indicated that

the concentration of leptin did not differ over the course of a year, again this may be the

result of lower specificity of the assay for emu leptin. If in the emu, leptin concentrations

increase when adiposity increases and suppress appetite, it would be impossible for the

emu to deposit large amounts of fat over the summer months as part of its annual cycle.

So, m y results suggest that leptin is not involved in appetite regulation in emus in the same

manner as it is in other species. The role of leptin in the control of appetite and adiposity

in the e m u will only be determined once effective methods for measuring leptin

concentrations in emu serum are developed. In the following chapters I will present

experiments aimed at improving the understanding of h o w appetite and adiposity are

controlled in the emu.

79

Chapter 4

The influence of dietary fat on food intake and adiposity

1. Introduction


3. Results

4. Discussion

Page

80

81

84

94

Chapter 4 - The influence of dietary fat on food intake and adiposity 80

1. Introduction

Seasonal fluctuations in feed intake and body weight are observed in emus and correlate

directly with their degree of adiposity (Mincham et al, 1998; Blache & Martin, 1999;

Blache et al, 2001b). Adiposity and appetite are decreased during the breeding season

and increased during the summer months. Decrease in appetite and adiposity during the

breeding season could be prevented by dietary interventions that disrupt energy balance.

For example, in chickens, feeding diets that are high in fat increases adiposity in

proportion to the diet's fat content (Edozien & Switzer, 1978; Deaton et al, 1981;

Boozer et al, 1995; Vila & Esteve-Garcia, 1996). In mammals, high fat diets also

induce hyperphagia, although this effect is variable between species and strains and

may be transient (Oscai et al, 1984; West et al, 1992; Hill et al, 1992; French et al,

1995; Smith et al, 1998). The variability of hyperphagia in response to high fat diets

may be due to the action of high fat diets to decrease the rate of stomach emptying

(Mateos et al, 1982). A decreased rate of stomach emptying would make the animal

feel sated for longer.

Diets high in fat disrupt energy balance by decreasing the metabolic rate and lowering

energy expenditure (Black et al, 1949; Storlien et al, 1986). The mechanism enabling

this could involve metabolic changes that can be assessed by measuring hormone

concentrations (Ramirez et al, 1990; Ahren & Scheurink, 1998). W h e n energy intake

exceeds the organism's requirements insulin rises, glucagon falls, and metabolism shifts

from catabolism of stored energy reserves to anabolism and storage of ingested energy

(Norman & Litwack, 1987). Depression of the thyroid hormones reduces the metabolic

rate and energy expenditure via depressed body temperature (Norman & Litwack,

1987).

This experiment was performed to determine if feeding diets that are high in fat to emus

during the summer months, when appetite is high, can further increase appetite and

adiposity. The following hypotheses were formulated:

1. Feeding emus a high fat diet for an extended period will increase adipose tissue

deposition and induce hyperphagia, and

2. Feeding emus a high fat diet will alter the concentration of metabolic hormones

to favour adipose deposition. This should result in higher concentrations of

insulin and lower concentrations of glucagon, triiodothyronine and thyroxine.



Experimental design

To test the hypotheses male emus were allocated into two groups to be fed either a high

(23.6%) or a low (4.5%) fat diet. Changes in appetite and adiposity were assessed by

measuring feed intake over the course of the experiment and performing carcass

composition analysis on the birds at the completion of the experiment. The

concentration of the insulin, glucagon, triiodothyronine and thyroxine were measured

over the course of the experiment by radioimmunoassay in order to determine if

metabolism had changed to favour adipose deposition.

Animals

Ten male emus (nine 3 year olds and one 4 year old) were penned individually, as

described in Chapter 2. They were allocated into two groups of similar average weights

(means ± s.e.m. - low fat diet group 41.5 ± 1.5 kg, high fat diet group 42.8 ±3.1 kg).

Treatments

The birds were offered diets differing in fat, but not protein content (Table 4.2). All

birds received a 4.5% fat diet (low fat) for a period of two weeks (Weeks -2 to 0). At

the end of the two weeks one group was given a 23.6% fat diet (high fat) for a period of

54 days. Due to difficulties with emus accepting changes in their feed, the switch to the

high fat diet was achieved by adding the high fat diet in increasing proportions to the

low fat diet, over the first two weeks (Weeks 1 and 2) of the treatment period.

Feeds were made up weekly or as required, with the addition of between 3 and 5%

water to the mix to aid in mixing and pelleting (Table 4.1). Flaked barley was reduced

to a powder using a grinding mill (Horwood Bagshaw Ltd) and the feed was

mechanically mixed in a Barrow Linton (Perth) mixer. Pellets were made with a 5 m m

diameter dye using a Lister pellet press. The size of the pellets approximates the size of

the commercial breeder pellet that the birds had previously been fed.


Table 4.1: Composition of the low and high fat diets.

Ingredient

Soya bean meal

Full-fat soya bean meal (Baye:

Full-fat canola seed (Davison

Barley flaked

Tricalcium phosphate

Limestone

Ethoxyquin

Vitamin E (50%)

Vitamin A/D (100%)

Sodium chloride

r XT-Soya)

Industries)

Brilliant blue food dye (Educational colours Pty Ltd)

low fat

45.3 kg

3.4 kg

5.2 kg

43.1 kg

1.5 kg

1.2 kg

4.96 g

4g

2g

141.1 g

200 ml

Diet (per 100 kg)

high fat

25.6 kg

9.1kg

52.7 kg

10.2 kg

1.5 kg

0.7 kg

25 g

20 g

2g

220 g

150 ml

Table 4.2: Macronutrient and energy content of the low and high fat diets.

Macronutrient

Fat(%)

Fibre (%)

Crude protein (%)

Total carbohydrate (%)

M E (MJ/kg)

E:P ratio

low fat

4.5

6.0

27.5

50.9

11.4

0.42

high fat

23.6

5.6

27.5

30.9

15.5

0.56

The carbohydrate content of the diet was not controlled and was higher in the low fat

diet. The fibre and total carbohydrate contents were estimated from data available for

components of the diet (McDonald et al, 1981).

Food intake

Food intake was recorded throughout the experiment, as described in Chapter 2, with

food and water offered ad libitum.

Live weight and blood sampling

O n Days -14, 0, 14, 28, 37 and 54 all birds were weighed and a 10 ml blood sample

collected, as described in Chapter 2.


Tissue collection and carcass composition

At the completion of the experiment (Day 56), birds were weighed and bled, prior to

being euthanased. Samples of liver and muscle were collected for analysis of fat

content. The empty carcass, subcutaneous and visceral fat depots, and the liver were

weighed to determine differences in adiposity. The methodology for these procedures

is given in Chapter 2.

Hormones and metabolites

Radioimmunoassays were performed for insulin, triiodothyronine, thyroxine and

glucagon for all samples as described in Chapter 2.


The carcass composition data for each tissue is expressed as a percentage of the live

weight of the animal. The significance of any differences between treatment groups,

were determined using Mann-Whitney U tests as described by Siegel (1956) using the

Statview™ 512+ program (1986). This statistical analysis was judged to be the most

appropriate method to analyse the data, as two independent groups were sampled from

and the sample size was small (n < 5). The significance of any differences within diets

over time, was determined using the Friedman two-way analysis of variance by ranks as

described by Siegel (1956). This statistical analysis was judged to be the most

appropriate as the samples were related and there were more than two time-points

involved. The results are indicated as the mean ± s.e.m.


3. Results

Feed Intake

There was a tendency for total feed and protein intakes to be lower on the high fat diet

(p = 0.075). The total amount of fat consumed was higher, and the total amount of fibre

consumed was lower in birds fed the high fat diet than in birds fed the low fat diet (p =

0.004, p = 0.048 respectively). Total energy consumption did not differ between the

two groups (Table 4.3), despite the higher energy content of the high fat diet (Table

4.2).

Table 4.3: Total feed, protein, fibre, fat and energy intakes over 68 days of male emus fed low and high

fat diets.

low fat diet high fat diet

Total feed intake (kg) 55.3 ±8.0 39.9 + 3.0

Total protein intake (kg) 15.2 + 2.2 11.0 + 0.8

Total fibre intake (kg) 3.34 ± 0.48* 2.30 ± 0.17*

Total fat intake (kg) 2.51 + 0.36* 6.50 ± 0.46*

Total energy consumption (MJ) 631 + 92 449 ± 40

Values are means ± s.e.m. A n * indicates significant difference (p < 0.05) between treatment groups.


For the majority of the experiment mean daily feed intake did not differ between the

groups. There were two exceptions with mean daily feed intake for Week 1 and Week 2

being lower for the high fat diet group (p = 0.008 and p = 0.028 respectively) (Fig. 4.1).

This period is also the transition period between the low fat and high fat diets and may

represent an initial rejection of the high fat diet. Within both the low and high fat diet

groups feed intake differed over time (p < 0.05 and p < 0.001 respectively).

Daily feed intake (kg)

1.6 |

1.4 I

1.2 I

1.0 I

0.8 I

0.6 '

0.4 I

0.2 I

0.0 ^ -2 -1 0 1 2. 3 4 5 4 7 8

Week

Figure 4.1: Average daily feed intake for each week of the experiment for male emus receiving either the low (open circle) or high fat diet (solid circle) for 8 weeks. The shaded area represents the transition

period of the high fat diet group from the low to the high fat diet. Values are means ± s.e.m. A n * indicates significant difference between treatment groups (p < 0.05).


Live weight

There was no difference in live weight between the high and low fat diet groups at any

time point and no effect of time on live weight within either group (Fig. 4.2).

Live weight (kg)

48'

46

44

42

40

38" r~ -14 14 28

- 1 —

42 56

Day

Figure 4.2: Mean live weight of male emus offered low (open circle) and high fat (solid circle) diets over a 54 day treatment period. The shaded area represents the transition period of the high fat diet group from the low to the high fat diet. Values are means ± s.e.m.


Carcass composition

There was no difference in the fat content of either muscle or liver between male emus

receiving either diet (Table 4.4).

Table 4.4: The fat content of muscle and liver tissue from male emus fed either low or high fat diets for 54 days.

Tissue low fat high fat

Muscle (g/kg of wet tissue) 4.93 ± 0.41 3.52 ± 0.90

Liver (g/kg of wet tissue) 28.3 ± 2.6 30.7 ± 1.6


The live weight at slaughter did not differ between treatment groups (Table 4.5). As

percentages of live weight, subcutaneous adipose, visceral adipose and total adipose

tissues, liver, empty carcass and testis weights did not differ between treatment groups

(Table 4.5). The percentage of digestive tract weight to live weight and digestive tract

length were greater in birds fed the high fat diet than in birds fed the low fat diet (p =

0.048 and p = 0.028 respectively) (Table 4.5).

Table 4.5: The body composition of male emus receiving either low or high fat diets for 54 days.

Parameter low fat high fat

Live weight at slaughter (kg) 42.6 ± 3.33 41.1+2.02

Subcutaneous adipose (%) 19.2 ± 1.33 17.5 ± 1.26

Visceral adipose (%) 7.4 ± 0.43 7.0 ± 0.34

Total adipose (%) 26.6 ± 1.42 24.5 ± 1.52

Liver (%) 1.4 ±0.1 1.2 ±0.05

Digestive tract weight (%) 1.5 ± 0.06 * 1.7 ± 0.11 *

Digestive tract length (%) 9.2 ± 0.28 * 11.2 ± 0.72 *

Testis (%) 0.014 ± 0.002 0.019 ± 0.003

Empty carcass (%) 52.3 ±1.12 54.4 ± 1.53

Values are means ± s.e.m. A n * indicates significant difference between treatment groups (p < 0.05). Parameters with a (%) next to them are given as the percentage tissue weight of live weight.



Serum glucagon did not differ between the high and low fat diet groups at any time

point (Fig. 4.3). Within each group there was no difference in glucagon concentration

over time.

Serum glucagon (pg/ml)

-14 0 14 28 42 56

Day

Figure 4.3: Serum glucagon concentrations of male emus offered low (open circle) and high (solid circle) fat diets for 54 days. The shaded area represents the transition period of the high fat diet group from the

low to the high fat diet. Values are means ± s.e.m.


There was a trend for insulin to decrease over time in both groups. However, this was

only significant for the high fat diet group (p < 0.05). The serum insulin concentrations

were lower in the birds receiving the high fat than the low fat diet at Day 14, Day 28

and Day 42 (p = 0.048, p = 0.016 and p = 0.048 respectively) (Fig. 4.4).

Serum insulin (ng/ml)

i 1 1 1 1 1

-14 0 14 28 42 56

Day

Figure 4.4: Serum insulin concentration of male emus offered low (open circle) and high (solid circle) fat diets foe 54 days. The shaded area represents the transition period of the high fat diet group from the low

to the high fat diet. Values are means ± s.e.m. A n * indicates significant difference between treatment

groups (p < 0.05).

2.0

1.8"

1.6"

1.4'

1.2"

1.0"

0.8"


In both groups it was observed that the insulin/glucagon ratio tended to decrease over

time. However, this trend only achieved significance for the high fat diet group (p <

0.05) (Fig. 4.5). The insulin/glucagon ratio tended to be lower in the high fat diet

group, however this was only significant on Day 28 (p = 0.032).

Plasma insulin/glucagon ratio

Day

Figure 4.5: Serum insulin/glucagon ratio of male emus offered low (open circle) and high (solid circle) fat diets for 54 days. The shaded area represents the transition period of the high fat diet group from the

low to the high fat diet. Values are means ± s.e.m. A n * indicates significant difference between

treatment groups (p < 0.05).

Chapter 4 - The influence of dietary fat on food intake and adiposity S

The serum triiodothyronine concentrations did not differ between the groups receiving

the high and low fat diets at any time point (Fig. 4.6). In both groups the

triiodothyronine concentration tended initially to decrease, however this trend was not

significant within either group over time.

Serum triiodothyronine (ng/ml)

1.6

1.4

1.2

1.0

0.8

0.6

0.4

0.2

0.0 - ' -14 0 14 28 42 56

Day

Figure 4.6: Serum triiodothyronine concentration in male emus offered low (open circle) and high (solid circle) fat diets for 54 days. The shaded area represents the transition period of the high fat diet group

from the low to the high fat diet. Values are means ± s.e.m.


The serum thyroxine concentration was higher in the high fat diet group at Day 14 and

lower at Day 42 (p = 0.048 and p = 0.028 respectively) (Fig. 4.7). Over time the

thyroxine concentration decreased in both the low fat and the high fat diet groups (p <

0.05 and p < 0.01 respectively).

Serum thyroxine (ng/ml)

25000 ]

20000 l

15000 j

10000

5000

0' -14 0 14 28 42 56

Day

Figure 4.7: Serum thyroxine concentration in male emus offered low (open circle) and high (solid circle) fat diets for 54 days. The shaded area represents the transition period of the high fat diet group from the low to the high fat diet. Values are means ± s.e.m. A n * indicates significant difference between treatment groups (p < 0.05).

Chapter 4 - The influence of dietary fat on food intake and adiposity 9'.

The triiodothyronine/thyroxine (T3/T4) ratio did not differ between the groups at any

time point (Fig. 4.8). There was no difference over time within the group receiving the

low fat diet. A difference in the T3/T4 ratio over time was observed within the group

receiving the high fat diet (p < 0.05).

T3/T4 ratio (1 x 10'4) 7.0

6.0

5.0

4.0

3.0

2.0

1.0

n J ' ' ' *—^- ~^ ' -14 0 14 28 42 56

Day

Figure 4.8: Serum T3/T4 ratio in male emus offered low (open circle) and high (solid circle) fat diets for 54 days. The shaded area represents the transition period of the high fat diet group from the low to the

high fat diet. Values are means ± s.e.m.

•;- . ;

; • •


4. Discussion

The results presented here do not support either of the hypotheses. Prolonged feeding

of a high fat diet did not increase adiposity or appetite, or alter hormone concentrations

to favour adipose storage (i.e. increased insulin, decreased glucagon, triiodothyronine

and thyroxine). The absence of increases in adiposity and appetite, and of changes in

hormone concentrations occurred despite more fat being consumed by the high fat diet

group. A s such, the absence of hormonal changes in these animals implies that in the

e m u this level of fat inclusion in the diet does not alter metabolism to favour energy

storage. The failure of the high fat diet to increase appetite and adiposity m a y be due to

the composition of the diet, taste aversions for components of the diet, or alterations in

digestive physiology in response to the high fat diet restricting feed intake.

Alternatively, the higher energy content of the high fat diet m a y have caused a

compensatory decrease in feed intake to maintain energy intake at a level appropriate to

the animal's requirements.

A factor that may have contributed to the failure of the high fat diet to increase

adiposity is its composition. First, the lower carbohydrate content of the high fat diet

m a y have resulted in a greater proportion of the ingested fat being oxidised to meet the

animal's energy requirements, particularly given the tendency for a lower level of

energy intake in the high fat diet group. Second, there is evidence that the type of fat

ingested can also affect the degree of adiposity. In rats it has been observed that

feeding isoenergetic diets containing either saturated or unsaturated fat results in greater

carcass adiposity in the saturated fat group (Shimomura et al, 1990). Broiler chickens

receiving isoenergetic diets of either saturated or unsaturated fat exhibited greater

abdominal adipose deposition in the group receiving saturated fat (Vila & Esteve-

Garcia, 1996; Sanz et al, 1999 & 2000). The difference between saturated fats and

unsaturated fats in their ability to induce adiposity is most likely related to differences

in their oxidation rates (Sanz et al, 2000). The greater the chain length of saturated fatty

acids the slower their rate of oxidation, a situation favouring storage (Leyton et al,

1987). The diets formulated for this experiment used full-fat canola and soyabean

meals as the primary sources of fat. As these fats are unsaturated, this m a y explain the

lack of difference in adiposity between the two groups. In chickens, at high levels of

inclusion, unsaturated fats, as fed in this experiment, increase adiposity (Vila & Esteve-


Garcia, 1996). As such, the high fat diet may not have contained enough unsaturated fat

to augment adipose deposition in emus.

An alteration in digestive function could also have been responsible for the lack of

effect of the high fat diet. Diets that are high in fat slow the rate of stomach emptying

(Mateos et al, 1982). It is therefore of interest that the weights and lengths of the

gastrointestinal tract, as proportions of live weight, were higher in the group receiving

the high fat diet. It is possible that this represents an adaptation to enable greater

absorption of dietary components. A slower rate of stomach emptying may also explain

the tendency for lower feed intake in the group receiving the high fat diet, as the animal

would feel sated for longer due to stomach distension. A slower rate of stomach

emptying in animals fed the high fat diet, and its satiating effects, may also explain the

failure of the diet to increase adiposity. A satiated animal eats less and a tendency for

lower energy intake was observed in the high fat diet group. As such, these animals

would expend more energy absorbing components from the diet, and the energy

obtained from this process is then more likely to be oxidised to meet the energetic

requirements of the animal. This situation would not favour adipose deposition.

The concentrations of insulin (lower), glucagon, triiodothyronine and thyroxine

(unchanged) also indicated that the metabolism of the birds fed the high fat diet did not

favour adipose deposition. As such, the hypothesis that metabolic changes that favour

adipose deposition will occur in response to a high fat diet can be rejected. However,

the hormonal data provides other insights into the metabolism of the e m u that m a y help

to explain the failure of the high fat diet to increase adiposity. One of these

explanations is that the e m u staunchly resists manipulations that alter metabolism. The

finding that significantly lower feed intake over a two week period was not associated

with significant changes in hormone concentrations supports this explanation. The

lower concentrations of insulin in the group receiving the high fat diet could mean that

they were in negative energy balance. The birds receiving the high fat diet also

consumed less food and had a lower energy intake, though neither was significant, that

would have made a state of negative energy balance possible. In addition, at the final

time-point the insulin concentration did not differ between the groups, following a two

week period of rising feed intake in the high fat diet group, indicating that the energy

balance had been restored in the high fat diet group. In contrast to insulin, glucagon did


not differ over the course of the experiment, implying that energy intake was sufficient.

The elevated concentration of glucagon at Day 14 was most probably due to the

decreased feed intake that occurred in response to the change in diet in the group

receiving the high fat diet. The decrease in feed intake observed represents voluntary

feed intake reduction in response to the presentation of an unfamiliar feed, a response

also observed by other researchers working with emus (Van Cleeff, 2000 personal

communication). Once accustomed to the new diet, feed intake was restored and the

concentration of glucagon in serum became comparable to that of birds receiving the

low fat diet. The insulin/glucagon ratio shows a strong tendency to be lower at all time

points following exposure to the high fat diet. This indicates that metabolism has been

altered to favour the utilisation of adipose stores rather than their deposition. The trends

observed in both groups for increasing glucagon and decreasing insulin over the course

of the experiment may reflect the onset of appetite suppression. This would suggest

that high fat diets do not sustain appetite in emus, in contrast to observations in mice

(Van Heek et al, 1997).

The rising feed intake of the group receiving the high fat diet during the final two weeks

of the experiment may explain the lack of difference in insulin between the two groups

at the final time point. The rising feed intake might also represent the induction of a

mechanism to sustain appetite when energy intake preceding the reproductive period is

insufficient. W h e n the feed intake of emus is restricted over summer (maximal

appetite) they respond by increasing their food intake and rate of weight gain in the

following months (O'Malley, 1999). The increased food intake and weight gain occurs

during a period when appetite would usually be declining. The ability to maintain their

appetite when the preceding period has not satisfied their energetic requirements is a

compensatory mechanism to ensure that the animals have sufficient reserves to survive

the breeding period. The initial voluntary reduction in feed intake observed in response

to the high fat diet and the lower feed intake while receiving the diet may have triggered

such a mechanism. Normally after periods of voluntary feed intake reduction, such as

occurs following stress in some birds (e.g. after handling), or after the completion of

incubation, appetite returns rapidly and lost weight is quickly restored (Van Cleeff,

2001; Zadworny et al, 1985). I did not observe a rapid compensatory appetite following

voluntary feed intake reduction in response to the high fat diet. This supports the idea


that the higher fat content decreased feed intake due to slower emptying of the stomach,

or as a result of a taste aversion to the feed.

The circulating concentration of triiodothyronine was not different at any time-point. In

both groups triiodothyronine concentrations initially decreased, and then increased from

Day 14, with this tendency strongest in the group receiving the low fat diet. The slower

rate of increase in the group receiving the high fat diet might reflect an effect of the

high fat diet to decrease the metabolic rate. It m a y also result from a compensatory

mechanism triggered by the voluntary starvation and lower energy intake of the group

receiving the high fat diet. The thyroxine concentration was higher in the group

receiving the high fat diet at Day 14 and lower at Day 42. The higher concentration of

thyroxine at Day 14 in the group receiving the high fat diet may be due to the voluntary

starvation observed in this group, as thyroxine is elevated in chickens in response to a

24 hour fast (Rosebrough & McMurtry, 2000). The lower concentration of thyroxine at

the end of the experiment in the group receiving the high fat diet may reflect an effect

of the high fat diet to decrease the metabolic rate. Thyroid hormone concentrations in

emus and ostriches are highly variable (Dawson, 1996; Dawson et al, 1996; Blache et

al, 2001c). In ostriches this variability is largely due to stress (Dawson & Deeming,

1997). The trends observed in triiodothyronine and thyroxine concentrations over the

course of the experiment may, therefore, be artefacts of the handling procedures

employed and the variability in thyroid hormone concentrations inherent in ratites. The

absence of any differences between groups in the ratio of T3/T4 supports this idea.

In conclusion, dietary fat, at the degree of inclusion used in this experiment, does not

increase adiposity, induce hyperphagia or cause a metabolic shift to favour adipose

deposition. There is evidence that unsaturated fats, as used in this experiment, do not

induce increases in adiposity as readily as saturated fats. If a higher level of fat

inclusion, or saturated fats had been used, adiposity m a y have increased. The greater

content of fat in the diet did cause changes in the digestive tract. These changes may

have compensated for a slower rate of stomach emptying, as observed in other species

receiving diets with a high fat content (Mateos et al, 1982). This slower rate of

emptying and possibly a taste aversion to the high fat diet m a y be responsible for the

lower feed intake and the lack of effect on adiposity in the group receiving this diet.

Additionally, the difficulty encountered in changing emus from one diet to another m a y


have confounded the experimental results. Ideally the experiment should have been

performed over a longer period. To overcome the limitations of dietary manipulations

in emus more direct mechanisms of stimulating appetite and altering metabolism could

be used. One such mechanism is treatment with glucocorticoids. In the following

chapter I have investigated the effect of dexamethasone, a synthetic glucocorticoid, on

appetite, adiposity and metabolism.

99

Chapter 5

The effect of dexamethasone on appetite, adiposity and the

hypothalamic expression of NPY in female emus.

Page

1. Introduction 100

Section A - Preliminary experiment

2A. Materials and Methods 103

3A. Results 105

4A. Discussion 106

Section B - Long-term dexamethasone treatment experiment

2B. Materials and Methods 107

3B. Results 110

4B. Discussion 122

Chapter 5 - The effect of dexamethasone on appetite, adiposity and 100 the hypothalamic expression of N P Y in female emus.

1. Introduction

The decrease in appetite observed in emus during the breeding season leads to

mobilisation of the adipose depot to meet energy demands (Blake, 1996; Blache &

Martin, 1999; Blache et al, 2001b). It is probable that changes in appetite are centrally

mediated, with the hypothalamus a logical site for investigation given its role in both

satiety and appetite.

There are a number of pathways that could be involved in the control of appetite and

adiposity in the emu. For example, pathways involving cocaine and amphetamine-

regulated transcript (CART), proopiomelanocortin ( P O M C ) , corticotrophin-releasing

hormone (CRH), orexin/hypocretin and agouti-related protein (AGRP), are involved in

the control of appetite and adiposity in other species (Mizuno et al, 1997; Ollmann et al,

1997; Sakurai et al, 1998; Heinrichs & Richard, 1999; Vrang et al, 1999). In this

experiment I have focused on the principal pathway involving N P Y as it is well

characterised for a range of species including several avian species.

Neuropeptide Y (NPY) is a potent, centrally acting, appetite stimulant (Levine &

Morley, 1984; Stanley & Leibowitz, 1984 & 1985; Stanley et al, 1985a & 1986;

Kuenzel et al, 1987; White, 1993; Woods et al, 1998; Inui, 1999a). It is possible that

mechanisms that disrupt appetite in the emu involve inhibition of N P Y . The regulation

of N P Y synthesis and secretion occurs at the level of the hypothalamic-pituitary-adrenal

(HPA) axis. This regulation occurs via changes in the concentration of insulin and

glucocorticoids, which act antagonistically (Strack et al, 1995). The regulation of N P Y

by insulin and glucocorticoids is a potential mechanism for the long-term regulation of

energy balance. As such, manipulating the hormonal balance could influence appetite

indirectly and overcome N P Y inhibition. The H P A axis is a logical target for

modification. Manipulation of the H P A axis to alter the regulation of N P Y could

maintain appetite and adiposity when appetite is normally diminishing. This

manipulation can be achieved with exogenous glucocorticoids, which diminish the

ability of leptin to depress appetite, increase the expression of N P Y m R N A and increase

the concentration of insulin (Campbell et al, 1966; Simon, 1984; Mercer et al, 1996;

Solano & Jacobson, 1999; Arvaniti et al, 1998; Zakrewska et al, 1997).


The action of glucocorticoids to first increase the expression of NPY mRNA, and

second diminish leptin's ability to increase appetite, promotes appetite. The elevated

concentration of insulin in response to glucocorticoid treatment increases lipogenesis,

gluconeogenesis and triglyceride deposition in adipose tissues. Increased appetite is not

consistently observed in animals receiving exogenous glucocorticoids. The absence of

increases in appetite m a y be due to the elevation of insulin inhibiting N P Y m R N A

expression, thereby removing the appetite stimulus (Schwartz et al, 1991; Dallman et

al, 1993). However, increased adiposity occurs with glucocorticoid treatment even

when appetite is not increased (Simon, 1984). As such, the elevated insulin

concentrations, and their promotion of adipogenesis, are primarily responsible for the

increased adiposity when appetite does not change in response to glucocorticoid

administration.

A synthetic glucocorticoid, dexamethasone, induces many of the changes mentioned

above in a wide range of species. In Syrian hamsters, chronic dexamethasone treatment

increases N P Y gene expression (Mercer et al, 1996). In rats, dexamethasone increases

both synthesis and release of N P Y (Corder et al, 1988). In sheep, injections of

dexamethasone increase appetite and reduce weight loss in animals exported live to the

Middle East (Adams & Sanders, 1992). Dexamethasone rapidly increases the

concentration of insulin and leptin in humans, and increases the concentration of insulin

in cattle (Corah et al, 1995; Kolaczynski et al, 1997; Miell et al, 1996; Papaspyrou-Rao

et al, 1997). This leads to insulin resistance and favours adipose deposition. The

elevation of leptin concentrations may also represent the induction of leptin resistance.

Glucocorticoid administration alters the metabolic rate. The administration of

glucocorticoids decreases the circulating concentration of thyroxine, while having

variable effects on the concentration of triiodothyronine (Darras et al, 1997).

Depression of the thyroid hormones, and subsequently metabolic rate, may contribute to

the increased adiposity observed in many species following dexamethasone treatment

(Walker & Romsos, 1993).

Glucocorticoids also influence reproductive function. In chickens, acute

dexamethasone treatment inhibits luteinising hormone secretion, rapidly reducing the

circulating luteinising hormone concentration (Wilson & Lacassagne, 1978). If the


reduction in luteinising hormone concentration is sustained by chronic treatment it

could act to maintain adiposity and appetite by removing the influence of reproductive

hormones on metabolism and appetite.

An experiment was performed to determine if dexamethasone treatment, during a period

when appetite is normally declining, enables appetite and adipose reserves to be

sustained. The following hypotheses were formulated:

1. Dexamethasone administration will increase the m R N A expression of N P Y ,

2. Long-term dexamethasone administration will decrease the concentrations of

triiodothyronine, thyroxine, and luteinising hormone, and increase the

concentrations of blood glucose and insulin, and finally

3. Increased expression of N P Y and metabolic changes induced by dexamethasone

will be associated with increased appetite and adiposity.

To enable these hypotheses to be tested it was necessary to determine the correct dosage

to use for emus. Therefore, a preliminary experiment was performed to determine the

correct dosage of dexamethasone to use in the chronic trial. Dexamethasone treatment

decreases the concentration of corticosterone to barely detectable levels (Buckland et al,

191 A; Etches, 1976; Corder et al, 1988; Corah et al, 1995; Zakrzewska et al, 1999). As

such, this can be used as an indicator of the onset and duration of the drug's effects.

The preliminary experiment is described in Section A. The long-term dexamethasone

experiment is described in Section B.


Section A - Preliminary experiment

2A. Materials and Methods

Experimental design

To determine the correct dosage of dexamethasone to use in the long-term experiment

male and female emus were treated with either a 6 m g or a 12 m g dose of

dexamethasone. Blood samples were collected at regular intervals and the

concentration of corticosterone was measured by radioimmunoassay to determine if the

doses used were capable of suppressing corticosterone production and the duration of

this effect.

Animals

Four male and four female emus that were approximately 12 months of age and hatched

in the same season were used for this experiment. The experiment was performed

during the non-breeding season. All the animals used for this experiment were in good

health.

Treatment

Half of the birds of each sex were implanted with 6 m g of dexamethasone. The

remaining birds were implanted with 12 m g of dexamethasone. The implants were

placed subcutaneously in the area behind the wing. To do this a small incision was

made in the skin and a purpose built tool was used to place the implant subcutaneously.

The implants were not removed after completion of the experiment.

Blood sampling

As described in Chapter 2, 10 ml blood samples were collected. Blood samples were

taken immediately before the implantation procedure, and again 7, 24, 72, 144, 240, 385

and 552 hours after implantation.


A radioimmunoassay was performed for corticosterone, as described in Chapter 2.


The significance of any differences between the 6 m g and 12 m g treated groups were

determined at each time-point by Mann-Whitney U tests as described by Siegel (1956)


using the Statview™ 512+ program (1986). This test was judged to be the most

appropriate as samples were taken from two independent groups and the sample size

was small (n = 4, n = 4). The significance of differences over time was determined

within groups by Friedman two-way analysis of variance by ranks as described by

Siegel (1956). This test was judged to be the most appropriate as the samples were

related and several time-points were under comparison. The results are indicated as the

mean ± s.e.m.

Chapter 5 - The effect of dexamethasone on appetite, adiposity and the hypothalamic expression of N P Y in female emus.

105

3A. Results


The serum corticosterone concentration did not differ between the dosages tested at any

time point. Over time both the 6 m g and 12 m g treated groups exhibited depression of

their corticosterone concentration in response to the dexamethasone implant (p < 0.01

and p < 0.05 respectively). The corticosterone concentration began increasing again

after 144 hours indicating the treatment duration was less than a week using implants of

dexamethasone (Fig. 5.1).

Serum corticosterone (ng/ul)

6 -i

4 -

3 -

0 100 200 300

Time (hours)

400 500 600

Figure 5.1: The serum corticosterone concentration of emus treated with a single subcutaneous dexamethasone implants of either 6 mg (solid circle) or 12 mg (open circle). Values are means ± s.e.m.


4A. Discussion

The preliminary experiment indicated that a single dose of dexamethasone between 6

and 12 m g decreased the corticosterone concentration, although the levels remained

detectable by the assay method employed. The results also indicate that using

subcutaneous implants the treatment was effective for approximately one week.

Trimedexil (dexamethasone trimethylacetate 5 mg/ml, Ilium Veterinary Products), was

chosen for use in the long-term dexamethasone treatment experiment. Trimedexil has

several advantages 1) it involves less handling stress as it is administered as a weekly

injection rather than weekly subcutaneous implants, and 2) it is a long-lasting liquid

preparation. This avoids problems relating to rapid clearance of the tablet form from

the circulation and also differences in the rate of release of the implants between birds

and over time. Both doses used in the preliminary experiment were equally capable of

depressing corticosterone and it was therefore decided to use a dose that fell between

those tested. As such, a dose of 10 mg/week was employed in the subsequent

experiment. There was no apparent difference in response to corticosterone between

males and females. Therefore, it was initially intended to include both males and

females in the long-term trial. However, once the experiment had commenced, three

birds in the male group were found to be females and another two died before the

experiment was completed. As a result there were not enough males to give meaningful

statistics and the male study group was abandoned. Prolonged treatment of catabolic

doses of dexamethasone can cause muscle wasting. To be certain that dexamethasone is

not catabolic when administered chronically at a dose of 10 mg/week, it is necessary to

determine if muscle wasting occurs. In the long-term trial this was assessed by

measuring the relative proportions of muscle and bone from a discrete region of the

carcass.


Section B - Long-term dexamethasone treatment experiment

2B. Materials and methods

Experimental design

To test the hypotheses outlined in the introduction adult female emus were allocated

into a control group receiving saline injections weekly or a treatment group receiving

dexamethasone injections weekly. The m R N A expression of N P Y was quantified in the

mediobasal hypothalamus/preoptic area at the completion of the treatment period by

ribonuclease protection assay. The concentration of triiodothyronine, thyroxine,

luteinising hormone and insulin were measured at regular intervals over the course of

the experiment by radioimmunoassay. Blood glucose was measured using a hand held

glucometer. The association between changes in N P Y , hormone and appetite and

adiposity, were assessed by measuring food intake over the course of the experiment,

weighing and fat scoring the animals at several time-points and performing carcass

composition analysis at the completion of the experiment.

Animals

Twelve female emus, aged between one and two years were used. The birds were

allocated into two groups of similar average weight. All birds were penned

individually, with commercial emu feed available ad libitum (Glen Forrest

Stockfeeders), see Chapter 2. The experiment commenced at the beginning of March,

coinciding with the start of the breeding season.

Treatments

The groups comprised dexamethasone treated (n = 7) and saline treated (n = 5) animals.

Treatments consisted of weekly injections of 10 m g of Trimedexil (dexamethasone

trimethylacetate 5 mg/ml, Ilium Veterinary Products) for six weeks, given as a single 2

ml intramuscular injection. Saline treated birds experienced the same handling and

received a 2 ml intramuscular injection of saline. The experiment was performed over a

nine week period consisting of pretreatment (Day -14 through to Day 0) and treatment

(Day 0 through to Day 49) periods.

Feed intake

Feed intake was recorded over the course of the experiment from Day -14. Recordings

were carried out on three days per week to give a weekly feed intake value, as described


in Chapter 2.

Live weight and fat scoring

Birds were weighed at three time-points over the course of the experiment (Day -14,

Day 14 and Day 49), immediately prior to bleeding and treatment. Fat scoring was

performed twice, at the commencement and completion of the experiment as described

in Chapter 2.

Blood sampling

As described in Chapter 2, 20 ml blood samples were collected weekly, immediately

prior to treatment. A n additional blood sample was collected from all birds on Day 2.

After the blood glucose measurement, the remaining blood was dispensed into separate

tubes for serum and plasma collection.

Tissue collection

At the completion of the experiment (Day 49), birds were bled and the blood glucose

concentration was recorded prior to the animals being euthanased with Lethabarb as

described in Chapter 2. The brain was removed and the hypothalamus and preoptic area

collected for R N A extraction as described in Chapter 2.

Carcass composition

Carcass composition analysis was performed as described in Chapter 2. The weights of

the subcutaneous adipose, visceral adipose and total adipose depots, and the liver, heart

and ovary were recorded for all birds. To assess the degree of muscle catabolism after

skinning and subcutaneous adipose tissue removal the total weight of the area between

the hip and knee was measured for the left leg. The individual contributions of muscle

and bone to this weight were then determined.


R N A was extracted as described in Chapter 2. The ribonuclease protection assay was

performed on total R N A extracted from the region containing the mediobasal

hypothalamus and the preoptic area, using the R P A III kit (Ambion) as described in

Chapter 2. The m R N A expression of N P Y and p-actin were determined using the N I H

Image program version 1.61. The sample number is one less for each group (ie


dexamethasone n=6, saline n=4), as a sample from each group was lost after collection.


Radioimmunoassay's were performed for corticosterone, insulin, triiodothyronine,

thyroxine, and luteinising hormone, as described in Chapter 2. immediately after each

bleed, measurements were made of blood glucose using a hand held blood glucose

meter as described in Chapter 2.


The expression of N P Y was determined as a value relative to the expression of P-actin

for each sample. The carcass composition data for each tissue was expressed as a

percentage of the live weight of the animal. For the ovary, the value was expressed as a

proportion of the ovary weight in grams, of the live weight in kilograms. Any effects of

dexamethasone to promote muscle catabolism were assessed by determining the

percentages of muscle and bone of the weight of the region between the hip and knee.

The significance of differences at each time-point was determined by Mann-Whitney U

tests as described by Siegel (1956) using the Statview™ 512+ program (1986). This test

was judged to be the most appropriate as two independent groups were sampled from,

the sample size was small (nl < 7, n2 < 5) and the number of birds in each group was

not equal. The significance of differences over time within groups was determined

using Friedman two-way analysis of variance by ranks as described by Siegel (1956).

This test was judged to be the most appropriate as the samples were related and more

than two time-points were involved. The Chi-Square test of independence (McGrath,

1997) was used to determine the significance of any differences in fat score observed

between groups at each time-point. This test was judged to be the most appropriate as

the data consisted of frequencies in discrete categories, and two independent groups

were under comparison. The Wilcoxon matched-pairs signed-ranks test (Siegel, 1956)

was used to determine the significance of any differences in fat score over time within

each group. This test was judged to be the most appropriate as the samples were related

and sample size was small (dexamethasone n = 4, saline n = 3). The results are

indicated as the mean ± s.e.m.


3B. Results


The expression of NPY mRNA did not differ between the dexamethasone treated (n =

6) and saline treated (n = 4) groups after 49 days of treatment (Fig. 5.2 and Table 5.1).

Table 5.1: The intensity of the mRNA expression of NPY relative to P-actin of female emus treated with either dexamethasone or saline for 49 days.

mRNA Dexamethasone Saline

(n = 6) (n = 4)

NPY/p-actin 0.51 ± 0.054 0.54 ± 0.054

Values are means ± s.e.m

Figure 5.2: The N P Y m R N A expression of female emus treated with either dexamethasone (lanes 1-6) or saline (lane 7-10) for 49 days. Lane 11 and 12 contain the P-actin and N P Y sense strand controls, respectively. Double-ended arrows indicate the position of the N P Y (a) and P-actin (b) bands in the samples. Single-headed arrows indicate the position of the N P Y (c) and P-actin (d) bands in the sense

strand controls.


Carcass composition

There was no difference in the weight of the subcutaneous or visceral adipose depots,

total adiposity, the heart or the hip to knee region as a percentage of live weight. The

weight of muscle and bone as a percentage of the hip to knee weight did not differ

between the groups (Table 5.2). The weight of liver as a percentage of live weight was

greater in the dexamethasone treated birds (p = 0.009). The proportion of ovary to live

weight was lower in the dexamethasone treated birds (p = 0.009) (Table 5.2).

Table 5.2: The carcass composition of female emus treated with either dexamethasone or saline for 49 days.

Parameter

Subcutaneous adipose (%)

Visceral adipose (%)

Total adipose (%)

Liver (%)

Ovary (g/kg lwt)

Heart (%)

Hip to knee (%)

Muscle (% of hip to knee wt)

Bone (% of hip to knee wt)

Values are means ± s.e.m. An :

Dexamethasone

(n = 7)

9.3 ± 1.47

5.7 ±0.73

15.0 ±1.98

2.18 ±0.124*

0.28 ±0.081*

1.09 ±0.059

15.9 ±0.74

76.2 ± 2.62

20.7 ± 0.88

Saline

(n = 5)

13.1 ±1.98

5.5 ± 0.43

18.6 ±2.35

1.35 ±0.085*

3.70 ±1.328*

1.00 ±0.043

15.4 ± 0.76

80.5 ±1.04

19.4 ±1.05

* indicates significant difference between groups (p = 0.009).


Feed intake

Feed intake was lower in the group treated with dexamethasone during W e e k 5 and

W e e k 6 (p = 0.024 and p = 0.024 respectively) (Fig. 5.3). Within groups feed intake

differed over time (a decreasing trend in the final three weeks) in the dexamethasone

treated group (p < 0.01). Over time feed intake did not differ in the saline treated group.

Average daily feed intake (kg)

1.0 1

0.9 j

0.8 ] -r

0.7 j -r

0.6 j f T i n "f W~}1 0.5 j I L I j IT

04 I I '' ' 1 I -' ; i i [

0.3 j j | ;

02 1 ! . I ^ i : i' • j! :

0.1 j

°-° TTTTTT Week

Figure 5.3: Average daily feed intake per week of female emus treated with either saline (black bar) or dexamethasone (grey bar) for 49 days. Values are means ± s.e.m. A n * indicates significant difference between groups (p = 0.024).


Live weight and fat scores

Live weight did not differ between groups at Day -14 or Day 14. At Day 49 live

weight was higher in the saline treated group (p = 0.024) (Fig. 5.4). Within groups the

live weight increased over time in the saline treated group (p = 0.039). Over time live

weight did not differ within the dexamethasone treated group.

Live weight (kg)

50

45

40

35

30

25

20

15

10

5

0 -14 14 49

Day

Figure 5.4: Live weight of female emus treated with either saline (black bar) or dexamethasone (grey bar) for 49 days, on Day -14, Day 14 and Day 49. Values are means ± s.e.m. An * indicates significant difference between groups (p = 0.024).

I_ JkX X


The fat score decreased in the group treated with dexamethasone after 49 days of

treatment (p = 0.023). The fat score showed a trend to increase in the group treated

with saline after 49 days of treatment (p = 0.05). However, the fat scores did not differ

on Day 0 or Day 49 between the treatment groups (Table 5.3).

Table 5.3: The fat scores of female emus on Day 0 and Day 49 of treatment with either dexamethasone or saline.

Day Dexamethasone Saline

(n = 7) (n = 5)

0 3.29 ±0.474 2.80 ±0.583

49 2.14 ±0.634 4.00 ±0.548




The serum insulin concentration did not differ between or within treatment groups at

any time point (Fig. 5.5).

2.8

2.6

2.4

2.2

2.0

1.8

1.6

1.4

1.2

1.0

0.8


•14 14 —r-

21

— i —

28 35 —r-42

— i

49

Day

Figure 5.5: Serum insulin concentration of female emus treated with either saline (open circle) or dexamethasone (solid circle) for 49 days. Values are means ± s.e.m.


The blood glucose concentration did not differ between the groups before treatment

commenced (Day -14, Day -7 and Day 0). Following the commencement of

dexamethasone treatment (Day 0), the blood glucose concentration was higher in the

group treated with dexamethasone (p < 0.003) (Fig. 5.6). Over time the blood glucose

concentration increased in the group treated with dexamethasone (p < 0.001). Over time

the blood glucose concentration decreased in the group treated with saline (p < 0.01).

Blood glucose (mmol/L)

15

14

13

12

11

10

9

8

7

6

5 -14 -7 0 7 14 21 28 35 42 49

Day

Figure 5.6: Blood glucose concentration of female emus treated with either saline (open circle) or dexamethasone (solid circle) for 49 days. Values are means ± s.e.m. A n * indicates significant difference

between groups (p < 0.01).


The serum thyroxine concentration was lower in the group treated with dexamethasone

than the group treated with saline from Day 7 until the completion of the experiment (p

= 0.001) (Fig 5.7). Within groups thyroxine decreased over time in the dexamethasone,

but not the saline treated group (p < 0.001).

Serum thyroxine (nmol/L)

i 1 1 1 1 1 1 1 1 i

-14 -7 0 7 14 21 28 35 42 49

Day

Figure 5.7: Serum thyroxine concentration of female emus treated with either saline (open circle) or dexamethasone (solid circle) for 49 days. Values are means ± s.e.m. A n * indicates significant difference between groups (p = 0.001).


The serum triiodothyronine concentration was higher in the dexamethasone than the

saline treated group on Day 7 and Day 14 (p = 0.009 and p = 0.015 respectively) (Fig.

5.8). Within the dexamethasone treated group triiodothyronine differed over time (p <

0.02). Over time there was no difference within the saline treated group.


2.0 '

1.8 '

1.6

1.4

1.2

1.0

0.8 '

0.6 '

0.4 '

0.2 '

0.0 -14 -7 0 7 14 21 28 35 42 49

Day

Figure 5.8: Serum triiodothyronine concentration of female emus treated with either saline (open circle) or dexamethasone (solid circle) for 49 days. Values are means ± s.e.m. An * indicates significant difference between groups (p < 0.05).


The triiodothyronine/thyroxine (T3/T4) ratio increased over time in the group treated

with dexamethasone, but did not change within the group treated with saline (p <

0.001). The T3/T4 ratio was higher in the group treated with dexamethasone than the

group treated with saline from Day 2 until the completion of the experiment (p < 0.05)

(Fig. 5.9).

T3/T4 ratio (1 x 10"3)

10.0 '

9-0 • *

8.0 '

7.0 '

-14 -7 0 7 14 21 28 35 42 49

Day

Figure 5.9: T3/T4 ratio of female emus treated with either saline (open circle) or dexamethasone (solid circle) for 49 days. Values are means ± s.e.m. An * indicates significant difference between groups (p <

0.05).


120

The serum luteinising hormone concentration was higher in the group treated with

dexamethasone than the group treated with saline at D a y 14 and D a y 21 (p = 0.037 and

p = O.OOlrespectively) (Fig. 5.10). There was no difference in the luteinising hormone

concentration over time within either group.

Serum luteinising hormone (ng/ml)

4.0

3.5

3.0

2.5

2.0

1.5

1.0

0.5

0.0 -14 14 21 28 35 42 49

Day

Figure 5.10: Serum luteinising hormone concentration of female emus treated with either saline (open circle) or dexamethasone (solid circle) for 49 days. Values are means ± s.e.m. An * indicates significant difference between groups (p < 0.05).


The serum corticosterone concentration of the group treated with dexamethasone was

higher at Day -14, and lower from Day 2 until the completion of the experiment (p =

0.04 and p < 0.02 respectively) (Fig. 5.11). Within groups the corticosterone

concentration decreased over time in the group treated with dexamethasone, but did not

change in the group treated with saline (p < 0.001).


14

12

10

8

6

4

2

0 -14 -7 0 7 14 21 28 35 42 49

Day

Figure 5.11: Serum corticosterone concentration of female emus treated with either saline (open circle) or dexamethasone (solid circle) for 49 days. Values are means ± s.e.m. A n * indicates significant

difference between groups (p < 0.05).


4B. Discussion

The results presented here do not support the hypotheses that prolonged treatment with

dexamethasone increases N P Y gene expression, or is associated with increased appetite

and adiposity. The hypothesis that dexamethasone treatment would change the

concentration of metabolic hormones is partially supported by the results, with

thyroxine decreased and blood glucose increased as predicted. The remaining

hormones measured did not change in the manner hypothesised. As N P Y is a potent

appetite stimulant the failure of dexamethasone to increase N P Y gene expression is

most likely responsible for the lack of increase in appetite. The failure to increase

adiposity m a y be due to the absence of both increased appetite and increased insulin

concentrations, and therefore adipogenesis. This implies that in the emu

dexamethasone treatment does not induce changes that facilitate increases in appetite or

adiposity. As such, long-term dexamethasone treatment is not a viable option for

maintaining or increasing adiposity in emus during periods when adipose stores are

mobilised.

Dexamethasone treatment also failed to depress triiodothyronine or luteinising

hormone. This could indicate that the dose used was too low. Muscle catabolism was

not observed following dexamethasone treatment, indicating that the dose of

dexamethasone given did not induce a catabolic state as is observed when administered

at high doses. The lower concentrations of corticosterone in the dexamethasone treated

birds from Day 2, indicates that the dosage did suppress the adrenocortical axis. It must

be noted that the significantly higher concentration of corticosterone prior to the

commencement of treatment (Day -14 and Day -7) in the dexamethasone treated group

resulted from the allocation process to obtain similar starting live-weights in both

groups. Three of the birds allocated to the dexamethasone treatment group were

extremely stressed by handling resulting in their corticosterone concentrations being

very high, and raising the group average. The thyroxine concentration was decreased

and the blood glucose concentration increased by dexamethasone treatment, as

hypothesized. However, as the thyroxine concentration decreased without increasing

adiposity it can be argued that decreased thyroxine was not associated with depression

of metabolic rate. The absence of any differences between the groups in their

triiodothyronine concentrations and adiposity despite significantly decreased thyroxine

and an increased T3/T4 ratio in the dexamethasone treated group indicates that


triiodothyronine may be more important than thyroxine in controlling metabolism in the

emu.

There was a tendency for the total and subcutaneous adiposity to be lower in the

dexamethasone treated group possibly due to the decreased feed intake observed in this

group over the final two weeks of the experiment. Dexamethasone treatment resulted in

an increased liver percentage of live-weight, most likely due to lipid infiltration.

Finally dexamethasone treatment was associated with suppression of ovarian

development. The depression of ovarian development may not have been a direct

effect, but rather a consequence of the suppressed appetite, weight loss and decreasing

adiposity during the final stage of the experiment in the dexamethasone treated group.

This is supported by the lack of any sustained differences in luteinising hormone

concentrations between the dexamethasone and saline treated groups over the course of

the experiment.

These results have two implications. First, that under these conditions NPY does not

play a major role in the control of appetite in the emu, as diminished appetite in the

dexamethasone treated birds at the completion of the experiment was not associated

with decreased N P Y gene expression. Indeed, it has been illustrated in mice that

multiple redundant pathways exist that control appetite in the absence of N P Y (Erickson

et al, 1996). Therefore, it is possible that genes other than N P Y could be of greater

importance to the control of appetite in emus under these conditions. Genes that could

be of greater importance than N P Y to the control of appetite in the emu include

orexin/hypocretin, proopiomelanocortin ( P O M C ) , agouti-related protein (AGRP),

cocaine and amphetamine-regulated transcript ( C A R T ) and corticotrophin-releasing

hormone (CRH). Second, as this experiment was performed during the period where

appetite is decreasing at the commencement of the breeding season, it is possible that

some breeding season or photoperiod-related factor was inhibiting the response of N P Y

to dexamethasone and also possibly its action to stimulate appetite.

There is evidence that the responsiveness of appetite to stimulating factors changes

seasonally in birds. One such factor that increases appetite via N P Y is prolactin. In

non-breeding ring doves prolactin increases appetite (Buntin & Figge, 1988; Hnasko &

Buntin, 1993; Buntin et al, 1999). In the ring dove prolactin increases appetite by


increasing NPY-immunoreactive cell bodies in the infundibular region of the

hypothalamus (Strader & Buntin, 2001). In contrast, prolactin administration to turkeys

during the reproductive period either decreases or has no effect on appetite (Denbow,

1986). As such, the diminished appetite that is observed during the breeding season in

the e m u could indicate inhibition of either NPY's response to prolactin, or of NPY's

ability to increase appetite. The inability of dexamethasone to increase N P Y m R N A

gene expression during this experiment could also be due to inhibition of NPY's

response to appetite stimulants via the action of photoperiod-related factors. The

potential for factors to inhibit NPY's response to appetite stimulants during the breeding

season will be investigated in the following chapter. I will investigate N P Y expression

during incubation in the male emu, a model that is characterized by elevated prolactin

concentrations.

125

Chapter 6

Changes in food intake, adiposity and the hypothalamic

expression of N P Y and VIP in response to incubation.

1. Introduction


3. Results

4. Discussion

Page

126

129

132

143

Chapter 6 - Changes in food intake, adiposity and the hypothalamic expression of N P Y and VIP in response to incubation

126

1. Introduction

Incubating male emus exhibit a much greater reduction in appetite and adiposity than their

non-incubating counterparts or females during the breeding season (Blake, 1996). The

decreased appetite of the incubating male is a form of anorexia. Naturally occurring

anorexia is observed in many species during activities such as, hibernation, incubation and

migration. This state of voluntary fasting occurs via a "sliding set point" for energy

reserves that allows the animal to maintain a depressed appetite, despite decreasing energy

reserves, during periods when appetite would hinder more important activities (Mrosovsky

& Sherry, 1980).

In almost all species studied, food restriction or starvation increases NPY gene expression

and corticosterone concentrations (Boswell et al, 1999; McShane et al, 1993; Chua et al,

1991; Dallman et al, 1999). In humans, increased N P Y gene expression is also observed

during anorexia nervosa (St0ving et al, 1999). There are a number of pathways that could

be involved in the control of appetite during incubation in the emu. For example,

pathways involving cocaine and amphetamine-regulated transcript (CART),

proopiomelanocortin ( P O M C ) , corticotrophin-releasing hormone (CRH),

orexin/hypocretin and agouti-related protein (AGRP), are involved in the control of

appetite and adiposity in other species (Mizuno et al, 1997; Ollmann et al, 1997; Sakurai et

al, 1998; Heinrichs & Richard, 1999; Vrang et al, 1999). In this experiment it was decided

to focus on the principal pathway involving N P Y as it is well characterised for a range of

species including several avian species. However, the expression of N P Y m R N A during

the period of voluntary starvation, which accompanies incubation in many avian species,

has not been investigated.

The onset of incubation behaviour in avian species is triggered by a rise in vasoactive

intestinal polypeptide (VIP) (Sharp et al, 1989). VIP stimulates prolactin release, inducing

and maintaining incubation behaviour (Chaiseha & El Halawani, 1999; El Halawani et al,

1996; Sharp et al, 1989). Prolactin has been shown to affect appetite, but its effects may

be dependent upon reproductive status. Prolactin increases appetite in ring doves (non-

breeding), and decreases appetite in laying turkeys (Denbow, 1986; Buntin & Figge, 1988;

Hnasko & Buntin, 1993; Buntin et al, 1999). In non-breeding ring doves, the central

administration of prolactin increases the number of NPY-immunoreactive cell bodies

Chapter 6 - Changes in food intake, adiposity and the hypothalamic 127 expression of N P Y and VIP in response to incubation

indicating that the stimulation of appetite by prolactin is mediated at least partially by NPY

(Strader & Buntin, 2001). During the breeding season, prolactin may be prevented from

affecting appetite by photoperiod-related factors. These photoperiod-related factors could

act to prevent prolactin's actions in several ways. First, in contrast to the non-breeding

birds, N P Y gene expression may not increase during the breeding period in response to

elevated prolactin concentrations (Strader & Buntin, 2001). Second, N P Y gene expression

might increase in response to elevated prolactin concentrations, but its ability to increase

appetite inhibited by other factors generated by the metabolic changes that promote energy

sparing mechanisms during incubation. One such factor could be corticotrophin-releasing

hormone (CRH). C R H acts centrally to inhibit the effects of N P Y on appetite (McCarthy

et al, 1993; Menzaghi et al, 1993). C R H also stimulates adrenocorticotrophin ( A C T H )

secretion, and this stimulates the release of glucocorticoids from the adrenal gland

(Kacsoh, 2000). Experimental treatments that increase C R H immunoreactivity or gene

expression in the hypothalamus, and the administration of C R H receptor agonists or C R H ,

are associated with increased concentrations of corticosterone (Gupta & Brush, 1998; Grill

et al, 2000; Bruijnzeel et al, 2001; Helmreich et al, 2001).

Metabolic changes may also be directly involved in the control of appetite during

incubation. In incubating birds increased glucagon and decreased insulin concentrations

promote lipolysis (Chieri et al, 1972; Cherel et al, 1988). The lipid fuels supplied by the

catabolism of adipose reserves decrease the utilisation of glucose and maintain a constant

blood glucose concentration (Groscolas & Rodriguez, 1981; Le M a h o et al, 1981).

Incubation is also associated with lower triiodothyronine and thyroxine concentrations,

which decrease metabolic rate and thereby spare energy reserves (Cherel et al, 1988;

Groscolas & Leloup, 1989). In emus, incubation is associated with increased glucagon and

decreased insulin and thyroxine concentrations (Van Cleeff, 2001). As such, the hormonal

balance of the incubating emu is consistent with a catabolic state. However, comparisons

of hormone concentrations between non-incubating and incubating male emus during the

breeding season have not been made. Also, differences in hormone concentrations

between non-incubating and incubating male emus may partially explain the greater degree

of appetite suppression in the incubating emu.

An experiment was designed to identify associations between metabolic hormone


concentrations and neuropeptides at the hypothalamic level, and the decrease in appetite

during incubation. The following hypotheses were formulated:

1. N P Y gene expression in the hypothalamus will be higher in incubating than in non-

incubating male emus,

2. Hypothalamic VIP gene expression and prolactin concentration will be higher in

incubating than in non-incubating male emus,

3. Hormonal changes consistent with a catabolic state (increased glucagon and decreased

insulin concentrations), and reduced resting metabolic rate (decreased thyroxine and

decreased or unchanged triiodothyronine concentrations) will be observed in

incubating emus, with blood glucose levels maintained at a constant level, and finally,

4. The concentration of corticosterone will be higher in the incubating than the non-

incubating male emus.



Experimental design

To test the hypotheses outlined above male emus were allocated into two groups, one

group was given eggs and allowed to incubate for 6 weeks. The second group received no

visual or tactile contact with eggs and constituted the control group. At the completion of

the experiment the region containing the mediobasal hypothalamus/preoptic area was

collected and R N A was extracted. The expression of m R N A for N P Y and VIP in the

mediobasal hypothalamus/preoptic area were quantified by ribonuclease protection assay.

In addition food intake was measured over the course of the experiment to determine if any

associations existed with the expression of N P Y m R N A . The concentration of prolactin,

insulin, glucagon, thyroxine, triiodothyronine and corticosterone were measured over the

course of the experiment by radioimmunoassay. The blood glucose concentration was

measured using a glucometer. In addition, carcass composition analysis was performed to

determine the extent of catabolism of adipose and selected muscle tissue.

Animals

Eleven sexually mature male emus were penned individually and given ad libitum access

to commercial breeder pellets (Glen Forrest Stockfeeds) as described in Chapter 2.

Treatments

From June, five birds from the incubating group were given an egg every two to three days

until they had nine eggs or commenced incubation. The eggs given to the birds were from

the previous breeding season and had been blown and refilled with sand to their original

weight to ensure that enough eggs were available for the experiment and prevent problems

with them going rotten. The eggs were then painted with a non-odorous paint

approximating the original shell colour to account for any leaching of colour that had

occurred. The birds were allowed to incubate for six weeks. A group of six birds were

used as the controls.

Feed intake

Feed intake was determined during W e e k 6 of incubation. A control bird was matched to

each incubator for feed intake measurement. The birds were slaughtered together at the

end of W e e k 6 of incubation so that effects of changing photoperiod on feed intake

Chapter 6 - Changes in food intake, adiposity and the hypothalamic " 130 expression of N P Y and VIP in response to incubation

between paired individuals were accounted for and differences in appetite observed could

be attributed to reproductive status.

Blood sampling

Once a week from W e e k 3 of incubation a 20 ml blood sample was collected, as described

in Chapter 2, from the incubator and its matched control.

Tissue collection

Following euthanasia the mediobasal hypothalamus and the preoptic area were excised as

described in Chapter 2 for R N A extraction and R P A analysis.

Carcass composition

The carcass composition was carried out as described in Chapter 2. The gastrocnemius

lateralis from the left leg of each bird was also dissected out and weighed to determine if

protein catabolism contributed to the loss of live weight observed in incubating birds.

RPA analysis

R N A was extracted as described in Chapter 2. The R P A was performed on total R N A

extracted from the region containing the mediobasal hypothalamus and the preoptic area,

using the R P A III kit (Ambion) as described in Chapter 2. The m R N A expression of N P Y ,

VIP and P-actin were determined by measuring the intensity of the bands for each gene

visualised on a 1 x T B E , 5 % acrylamide gel. The band intensity was determined using the

N I H Image program version 1.61.


Radioimmunoassays were performed for insulin, triiodothyronine, thyroxine, prolactin,

corticosterone and glucagon for all samples as described in Chapter 2. Immediately after

each bleed measurements were made of blood glucose using a hand held blood glucose

meter (Precision Q.I.D, Medisense) as described in Chapter 2.


The expression of VIP and N P Y were determined as values relative to the expression of (3-

actin for each sample. For the carcass composition analyses each tissue weight was


converted to a percentage of the live weight of the animal. The significance of any

differences observed between incubating and non-incubating male emus was determined at

each time-point using Mann-Whitney U tests as described by Siegel (1956) using the

Statview™ 512+ program (1986). The significance of trends over time within each group

was determined using the Friedman two-way analysis of variance by ranks as described by

Siegel (1956). Non-parametric statistical analysis was judged to be the most appropriate as

two independent groups were sampled from, the sample size was small, and the number of

birds in each group was not equal. The results are indicated as the mean ± s.e.m.

Chapter 6 - Changes in food intake, adiposity and the hypothalamic

expression of N P Y and VIP in response to incubation

132

3. Results

RPA

The level of VIP m R N A expression was higher in the incubating males and they also

displayed a trend for elevated N P Y m R N A expression (p = 0.026 and p = 0.165

respectively) (Fig. 6.1 and Table 6.1).

Table 6.1: The m R N A expression of N P Y and VIP relative to P-actin of incubating (n=5) and non-

incubating (n=6) male emus.

mRNA Incubators Non-incubators

NPY/|3-actin

VEP/p-actin

0.69 ± 0.046

0.62 ± 0.046*

0.53 ± 0.083

0.47 ± 0.055*

Values are means ± s.e.m. A n * indicates significant difference between groups (p = 0.026).

Figure 6 1- The N P Y and VIP m R N A expression of incubating (lanes 1-5) and non-incubating (lanes 6-11) male emus determined using the R P A technique. Lanesl2, 13 and 14 contain respectively, the sense strand

positive controls for emu p-actin, N P Y and VIP. Arrows indicate the position of the bands for VIP (a, d),

N P Y (b, e) and P-actin (c, f) from samples and sense strand controls respectively.


Food intake, carcass composition and live weight

Food intake, live weight and percentage total fat, subcutaneous fat and visceral fat weights

of live weight and the percentage liver weight of live weight were all lower in the

incubating group (p = 0.002, p = 0.015, p = 0.009, p = 0.015, p = 0.041 and p = 0.041

respectively) (Table 6.2). The percentage of the gastrocnemius lateralis muscle of live

weight was higher in the incubating group (p = 0.026) (Table 6.2).

Table 6.2: The total feed intake over Week 6 of incubation and the body composition of incubating and non-

incubating male emus at the end of Week 6 of incubation.

Parameter Incubators Non-incubators

Total feed intake (kg) 0.20 ± 0.148* 6.93 ± 1.581*

Live weight (kg) 35.9 ± 1.95* 41.8 ± 1.03*

Subcutaneous adipose (%) 11.7 ±2.14* 17.6 ±1.11*

Visceral adipose (%) 4.09 ±0.889* 5.93 ±0.518*

Total adipose (%) 15.8 ± 2.91* 23.5 ± 1.24*

Liver (%) 1.17 ±0.152* 1.74 ±0.152*

Gastrocnemius lateralis muscle (%) 1.53 ± 0.065* 1.37 ± 0.042*

Values are means ± s.e.m. An * indicates significant difference between groups (p < 0.05).



The serum prolactin concentration was higher in the incubating males during Week 3,

Week 4, Week 5 and Week 6 (p = 0.004, p = 0.048, p = 0.002 and p = 0.002 respectively).

Within groups there were no difference in prolactin concentration over time (Fig. 6.2).

Serum prolactin (ng/ml)

14

12

10

8

6

4

2

2 3 4 5 6 7

Week

Figure 6.2: The serum prolactin concentration from Week 3 to Week 6 of incubation of incubating (solid circle) and non-incubating (open circle) male emus. Values are means ± s.e.m. An * indicates significant difference between groups (p < 0.05).

T 1 1 r


The blood glucose concentration did not differ between or within either group at any time

point (Fig. 6.3).

Blood glucose (mmol/L)

8.5

8.0

7.5

7.0

6.5

6.0 2 3 4 5 6 7

Week

Figure 6.3: Mean blood glucose concentrations from Week 3 to Week 6 of incubation of incubating (solid circle) and non-incubating (open circle) male emus. Values are means ± s.e.m.


There was a tendency for the serum glucagon concentration to be higher at all time points

in the incubating group. Serum glucagon concentrations were higher in the incubating

group, at W e e k 4 and W e e k 5 of incubation (p = 0.028 and p = 0.032 respectively). Within

groups glucagon concentrations did not differ over time (Fig. 6.4).

Plasma glucagon (pg/ml)

250

200

150

100

50

2 3 4 5 6 7

Week

Figure 6.4: Mean serum glucagon concentrations from Week 3 to Week 6 of incubation of incubating (solid circle) and non-incubating (open circle) male emus. Values are means ± s.e.m. A n * indicates significant


T 1 1 1 1


137

The serum insulin concentration did not differ between or within groups at any time point

(Fig. 6.5).


1.4

1.3

1.2

1.1

1.0

0.9

0.8

0.7

0.6

Week

Figure 6.5: Mean serum insulin concentrations from Week 3 to Week 6 of incubation of incubating (solid circle) and non-incubating (open circle) male emus. Values are means ± s.e.m.


138

The insulin/glucagon ratio was higher in the non-incubating males at W e e k 4 and W e e k 5

of incubation (p = 0.028 and p = 0.032 respectively). There was also a tendency for the

ratio to be higher in the non-incubating males at W e e k 3 (p = 0.057). Within groups the

ratio did not differ over time for either group (Fig. 6.6).

Insulin/Glucagon ratio

45

40

35

30

25

20

15

10

5

4 5

Week

Figure 6.6: Mean serum insulin/glucagon ratio from Week 3 to Week 6 of incubation of incubating (solid circle) and non-incubating (open circle) male emus. Values are means ± s.e.m. An * indicates significant difference between groups (p < 0.05).


The serum triiodothyronine concentration did not differ between or within groups at any

time point (Fig 6.7).


0.6

0.5

0.4

0.3

0.2

0.1

°-° 2 3 4

Week

Figure 6.7: Mean serum triiodothyronine concentrations from Week 3 to Week 6 of incubation of incubating (solid circle) and non-incubating (open circle) male emus. Values are means ± s.e.m.


The serum thyroxine concentration was higher in the non-incubating males at each

sampling point (p = 0.004) (Fig. 6.8). Within groups there was an effect of time in the

incubating group (p < 0.05). Over time the thyroxine concentration did not change in the

non-incubating group.

Serum thyroxine (nM/L)

5000 -

4000

3000

2000

1000

0' 2 3 4 5 6 7

Week

Figure 6.8: Mean serum thyroxine concentrations from Week 3 to Week 6 of incubation of incubating (solid circle) and non-incubating (open circle) male emus. Values are means ± s.e.m. An * indicates significant


f

Chapter 6 - Changes in food intake, adiposity and the hypothalamic 141 expression of NPY and VIP in response to incubation

The triiodothyronine/thyroxine (T3/T4) ratio did not differ between or within groups at any

time point (Fig. 6.9).

T3/T4 ratio (1 x 10"4)

14.0 "

12.0 "

10.0 "

8.0 '

6.0 "

4.0 "

2.0 '

0 ' 2 3 4 5 6 7

Week

Figure 6.9: The T3/T4 ratio from Week 3 to Week 6 of incubation of incubating (solid circle) and non-incubating (open circle) male emus. Values are means ± s.e.m.


The serum corticosterone concentration tended to be higher at all time points in the

incubating emus (Fig. 6.10). This trend was significant at Week 3 and Week 6 (p = 0.028

and p = 0.048 respectively). Within groups the concentration of corticosterone did not

differ over time for either group.


2.5 "

2.0 "

1.5

1.0 '

0.5

2 3 4 5 6 7

Week

Figure 6.10: Mean serum corticosterone concentrations from Week 3 to Week 6 of incubation of incubating (solid circle) and non-incubating (open circle) male emus. Values are means ± s.e.m. A n * indicates

significant difference between groups (p < 0.05).


4. Discussion

The hypothesis that hypothalamic N P Y gene expression is higher in incubating than non-

incubating emus is not supported by the results, although there was a tendency for higher

levels of expression in the incubating group. The results support the hypothesis that

hypothalamic VIP expression and circulating prolactin levels are increased in incubating

emus. This suggests that as with other avian species, in emus, VIP and prolactin are

involved in the maintenance of incubation behaviour (Mauro et al, 1989; Sharp et al,

1989). The hypothesis that emus enter a catabolic state during incubation, with increased

glucagon and thyroxine concentrations and a tendency for decreased insulin concentrations

is supported by the results. The hypothesis that corticosterone concentration is higher in

incubating emus is also supported, with a tendency for higher concentrations in the

incubators throughout the study period and significantly higher concentrations at two of

these time points.

The absence of a significant difference in NPY expression between incubating and non-

incubating emus during the breeding season raises some interesting possibilities. First,

N P Y expression may be at high levels in both incubators and non-incubators during the

breeding period due to diminishing adipose reserves. If N P Y expression is high, the

diminished appetite during the breeding period is probably the result of inhibition of

appetite promotion by N P Y . A candidate for the role of N P Y inhibitor is corticotrophin-

releasing hormone (CRH) a centrally acting hormone that prevents N P Y increasing

appetite (McCarthy et al, 1993; Menzaghi et al, 1993). The higher concentration of

corticosterone in the incubating birds lends support to a role for C R H , as C R H

administration, increased C R H expression or immunoreactivity, and C R H receptor

stimulation are associated with increased corticosterone concentration (Gupta & Brush,

1998; Grill et al, 2000; Bruijnzeel et al, 2001; Helmreich et al, 2001). Alternatively, the

higher concentration of corticosterone in the incubating emus in this experiment may

reflect greater stress due to the animals being in negative energy balance. Second, the

ability of N P Y to respond to decreasing adipose reserves may be inhibited during the

breeding season. Higher concentrations of prolactin in the incubating birds were not

associated with elevations in the level of N P Y m R N A expression. This result is in contrast

to the finding that N P Y was elevated in ring doves administered prolactin during the non-

breeding period (Strader & Buntin, 2001). The absence of significant differences in N P Y


mRNA expression supports the idea that NPY's ability to respond to factors, such as

prolactin, that would normally stimulate N P Y expression and thereby appetite, could be

inhibited during the breeding season. A final possibility is that N P Y may not have a

pivotal role in the regulation of body reserves in the emu. Appetite in emus might be

controlled at the hypothalamic level by other appetite mediators, such as, orexin,

proopiomelanocortin ( P O M C ) , agouti-related protein (AGRP), C R H or cocaine and

amphetamine-regulated transcript (CART).

As expected the anorexia and subsequent negative energy balance during incubation led to

catabolism of adipose and liver energy reserves to meet the animals energetic

requirements, resulting in lower proportions of adipose and liver tissue to live weight. The

catabolic state did not include depletion of muscle glycogen or protein as evidenced by the

higher proportion of the gastrocnemius muscle to live weight in the incubating birds. This

does not mean that muscle was increased during incubation, rather that muscle mass was

unchanged and subsequently increased as a proportion of live weight as the adipose stores

and live weight decreased.

The hormonal data also supports a catabolic state in the incubators whose glucagon

concentrations were higher, insulin concentrations tended to be lower and ratio of insulin

to glucagon was lower. As expected these changes in insulin and glucagon maintained a

constant level of blood glucose that did not differ from that of the non-incubating birds.

The expectation that the decreased activity and feed intake during incubation would lower

the metabolic rate is supported by the lower concentration of thyroxine in the incubators.

However, triiodothyronine and the T3/T4 ratio were not different between the two groups.

Overall these results suggest that incubation in the emu is associated with a decrease in the

metabolic rate and catabolism of adipose reserves. Other researchers have also observed a

decreased metabolic rate in the emu during incubation (Buttemer & Dawson, 1989a &

1989b).

As ad libitum food was available, the low level of intake of the non-incubating, and the

anorexia of the incubating birds, resulted from depressed appetite. In the following chapter

I examined the response of N P Y and VIP m R N A expression and the concentration of

metabolic hormones to short-term starvation in order to obtain a better understanding of


the control of appetite and adiposity in the emu. Imposed starvation induces many of the

alterations in metabolic hormones that are observed during anorexia. However, imposed

starvation is more stressful than naturally occurring anorexia and differs significantly in

that it is associated with increased appetite. The increase in appetite is mediated by

increased N P Y expression and changes in metabolic hormones. As such, investigating

appetite and adiposity using the model of short-term starvation will provide further insights

into the control of appetite and adiposity in the emu.

146

Chapter 7

The effect of short-term starvation on the hypothalamic

expression of NPYand VIP during a period of elevated

appetite.

1. Introduction


3. Results

4. Discussion

Page

147

149

152

159

Chapter 7 - The effect of short-term starvation on the hypothalamic, 147 expression of N P Y and VIP during a period of elevated appetite

1. Introduction

Throughout the course of the year emus undergo large shifts in their food intake, body

weight and metabolism. These changes include anorexia in incubating males coupled with

extensive utilisation of the adipose stores. Following the cessation of the breeding period

appetite is up-regulated and fat is deposited.

In Chapter 6, incubation was associated with increased expression of VIP mRNA in the

mediobasal hypothalamus/preoptic area, and a tendency towards elevated N P Y m R N A

expression. In Chapter 5, N P Y m R N A expression was unchanged in the same region by

dexamethasone treatment despite decreased appetite and a tendency for lower levels of

adiposity in the treated birds. As Chapters 5 and 6 investigated birds during the breeding

season, the actions and responses of N P Y to decreasing adiposity may have been

suppressed by factors associated with reproduction and photoperiod.

There are a number of pathways that could be involved in the control of appetite and

adiposity in the emu. For example, pathways involving cocaine and amphetamine-

regulated transcript (CART), proopiomelanocortin ( P O M C ) , corticotrophin-releasing

hormone (CRH), orexin/hypocretin and agouti-related protein (AGRP), are involved in the

control of appetite and adiposity in other species (Mizuno et al, 1997; Ollmann et al, 1997;

Sakurai et al, 1998; Heinrichs & Richard, 1999; Vrang et al, 1999). In this experiment it

was decided to focus on the principal pathway involving N P Y as it is well characterised for

a range of species including several avian species. To clarify the role of N P Y , and

investigate if VIP has a role in the control of appetite in emus, it is necessary to determine

how they respond to imposed starvation when appetite is maximal. In chickens, N P Y

m R N A expression is increased by starvation or feed restriction (Boswell et al, 1999). The

effects of starvation on VIP at the brain level have not been widely studied. However, in

rats the concentration of VIP in the brain is not altered by starvation (Shulkes et al, 1983).

Starvation, in addition to its central effects, alters the metabolic status of the animal. When

energy intake is insufficient animals enter a catabolic state that, in birds, is characterised

by decreased insulin and increased glucagon (Sitbon & Mialhe, 1978; Rosebrough et al,

1984). Insufficient energy intake leads to the catabolism of adipose and protein reserves to

meet the animals continuing energy requirements. Catabolism of adipose tissue increases

the concentration of P-hydroxybutyrate, while protein breakdown increases the


concentration of urea (Le M a h o et al, 1981; Garcia-Rodriguez et al, 1987). In chickens, a

two day fast decreases serum triiodothyronine and increases serum thyroxine

concentrations (Van der Geyten et al, 1999). Corticosterone concentrations are also

elevated by starvation and are associated with increases in N P Y expression (Dallman et al,

1999; Geris et al, 1999). In addition, adrenalectomy of rats decreases N P Y elicited food

intake by 60-71% and this is returned to unoperated levels by corticosterone replacement

(Stanley et al, 1989). It can be inferred from this that corticosterone is required for

exogenous N P Y to affect appetite. The rise in N P Y expression in response to starvation

follows the increase in corticosterone (Dallman et al, 1999). As such, increased

corticosterone may be a requirement for increases in N P Y expression or for potentiation of

its effects on appetite.

This experiment was performed to determine how short-term starvation during a period of

elevated appetite affects metabolism and the m R N A expression of N P Y and VIP. The

following hypotheses were tested:

1. Starvation, when appetite is high, will increase the expression of N P Y m R N A without

affecting VIP m R N A expression in the mediobasal hypothalamus/preoptic area,

2. That starvation would alter the concentration of hormones and metabolites to favour

catabolism of adipose tissue in the starved birds (concentrations of glucagon, (3-

hydroxybutyrate and thyroxine will increase and insulin and triiodothyronine will

decrease). As the starvation period is short, catabolism of protein is not expected to be

initiated, therefore the concentration of urea will not change, and finally

3. Elevated corticosterone concentrations will be observed in response to starvation.

Chapter 7 - The effect of short-term starvation on the hypothalamic, expression of N P Y and VIP during a period of elevated appetite

149


Experimental design

To test the hypotheses outlined above male and female emus were allocated into two

groups. The first group was used as a control, while the second group was subjected to a

one week period of starvation. At the completion of the starvation period the region

containing the mediobasal hypothalamus/preoptic area was collected and R N A was

extracted. The expression of N P Y and VIP m R N A in the mediobasal

hypothalamus/preoptic area was quantified by ribonuclease protection assay. The

concentrations of insulin, glucagon, thyroxine, triiodothyronine and corticosterone were

measured before and at the completion of the treatment period by radioimmunoassay. The

concentration of urea and (3-hydroxybutyrate were measured before and at the completion

of the treatment period by enzymatic and spectrophotometric assays respectively. In

addition, the birds were weighed before, and at the completion of the treatment period, and

carcass composition analysis was performed on a subset of birds to determine the extent to

which the adipose reserves were catabolized (Fig. 7.1).

Day

Bleed and

T i 1

-14

weigh

Pre-treatment

Feed intake both groups

I I I I -7 0

Treatment I I Control feed intake

Bleed and weigh

I

7

Figure 7.1: The sampling and measurement schedule for the pre-treatment and treatment periods.

Animals

A group of sixteen emus (seven males and nine females) were divided into two groups

comprising eight birds starved (five female and three male) and eight control birds (four

female and four male). All animals were kept in individual pens.

Treatment

The experiment commenced mid-November, a period when appetite is maximal. All the

birds were given ad libitum access to a commercial pellet feed (Glen Forrest Stockfeeders)


for one week. The feed was then removed from the eight animals in the starved group for

a period of seven days.

Measurement of feed intake

Feed intake was determined for individual birds for one week during the pre-treatment

period in both groups and for the control group over the course of the treatment period

(Fig. 7.1).

Sample collection

The birds were weighed and 20 ml samples of blood were collected at the start of the pre-

treatment period. At the completion of the treatment period, 20 ml blood samples were

collected and the birds were euthanased with Lethabarb as described in Chapter 2 (Fig.

7.1). The brains were removed from a subset of the animals (four control males, three

starved males and one starved female), and the mediobasal hypothalamus and preoptic area

tissues obtained and stored for R N A extraction and R P A analysis as described in Chapter

2.

Carcass composition

The carcass was weighed. The subcutaneous and visceral fat depots and the liver were

removed and weighed to determine differences between the groups in adiposity for a

subset of the animals used (same subset used for R P A analysis).

RPA analysis

R P A analysis was performed on the subset of samples collected. R N A was extracted as

described in Chapter 2. The R P A was performed on total R N A extracted from the region

containing the mediobasal hypothalamus and the preoptic area, using the R P A III kit

(Ambion) as described in Chapter 2.


Radioimmunoassays were performed for corticosterone, insulin, triiodothyronine,

thyroxine and glucagon for all samples as described in Chapter 2. 6-hydroxybutyrate and

urea assays were performed by the Agriculture Western Australia Animal Health

Laboratory. P-hydroxybutyrate was measured using a spectrophotometric assay

(McMurray et al, 1984). Urea was measured using the enzymatic U V test,


urease/glutamate dehydrogenase ( G L D H ) method. The urease/GLDH method incorporated

assays for urea, using the Trace Urea (Urea Nitrogen) Reagent Kit (Roche), and G L D H ,

using the G L D H M P R 1 Reagent Kit (Roche). Both assays were performed on a Roche

Cobas Mira S Automated Chemistry Unit. All samples for each metabolite were run in a

single assay.


Before the starvation period commenced (pre-treatment) the birds were weighed and blood

samples were collected and feed intake monitored, to enable within group comparisons to

be made over time (Fig. 7.1). The expression of VIP and N P Y were determined as values

relative to the expression of B-actin for each sample. The carcass composition data for

each tissue is expressed as a percentage of the live weight of the animal. The nature of

differences between the starved and control groups was determined using Mann-Whitney

U tests as described by Siegel (1956). The Wilcoxon matched-pairs signed-ranks test

(Siegel, 1956) was used to determine differences within groups over time using the

Statview™ 512+ program (1986). Non-parametric statistical analysis was judged to be the

most appropriate as two independent groups were sampled from, the sample size was

small, and the number of birds in each group was not equal. The results are indicated as

the mean ± s.e.m.


3. Results

RPA

There was no difference between control and starved male emus in the level of expression

of either VIP or N P Y after one week of starvation (p = 0.343 and p = 0.100 respectively)

(Table 7.1).

Table 7.1: The mRNA expression of NPY and VIP relative to 8-actin of starved and control male emus.

~~ m R N A ~ Control Starved

NPY/8-actin 1.07 ±0.107 0.77 ±0.168

VIP/6-actin 1.25 ±0.284 1.82 ±0.321


Feed intake

Feed intake did not differ between the groups for either sex in the pre-treatment period.

There was no change in feed intake of the control males over time, however the feed intake

of the females was higher during the treatment period (p = 0.0336) (Table 7.2).

Table 7.2: The average daily feed intake of male and female emus in control and starved groups one week before, and during a subsequent one-week treatment period.

pre-treatment

treatment

Control female

(kg/day)

(N=4)

0.45±0.07*

1.00±0.04*

Starved female

(kg/day)

(N=5)

0.55±0.20

-

Control male

(kg/day)

(N=4)

0.60±0.17

0.85±0.22

Starved male

(kg/day)

(N=3)

0.73±0.13

-

Values are means ± s.e.m. An * indicates significant difference within groups over time (p = 0.0336).


Live weight

Live weight did not differ between the groups at the start of the pre-treatment period or at

the completion of the treatment period. There was no change in live weight of the control

or starved males over time. However, live weight increased in the control and decreased in

the starved females over time (p = 0.0336 and p = 0.0217 respectively) (Table 7.3).

Table 7.3: The live weight of male and female emus in the control and starved group before and after a 7 day treatment period.

pre-treatment

treatment

Control female (kg)

(N=4)

51.0 + 2.1*

53.6 ±2.7*

Starved female (kg)

(N=5)

52.4 ± 2.5"

50.9 ± 2.5"

Control male (kg)

(N=4)

49.5 ±1.4

49.3 ±1.6

Starved male (kg)

(N=3)

47.2 ±3.1

46.0 ± 3.3

Values are means ± s.e.m. The * and a indicate significant difference within groups over time (p < 0.05).

Carcass composition

The visceral, subcutaneous and total fat as a percentage of live weight did not differ

between the groups. Liver weight as a percentage of live weight was higher in the control

group at the completion of the treatment period (p = 0.029) (Table 7.4).

Table 7.4: The body composition of control and one week starved male emus.

Parameter Control Starved

Subcutaneous adipose (%) 22.6 ±1.09 22.8 ±1.00

Visceral adipose (%) 9.90 ± 0.850 8.93 ± 0.869

Total adipose (%) 32.6 ± 0.33 31.8 ± 1.56

Liver (%) 1.59 ± 0.092* 1.08 ± 0.150*

Values are means ± s.e.m. A n * indicates significant difference between groups (p = 0.029)



The insulin concentration was lower in the female starved group at the completion of the

treatment period (p = 0.008), but not pre-treatment. The insulin concentration did not

differ between the male groups at either time point. Within groups the insulin

concentration decreased over time in the starved females (p = 0.022), but not the starved

males or the control groups. There was a tendency for the insulin concentration to increase

over time in the control female group (p = 0.072) (Table 7.5).

Table 7.5: Serum insulin concentrations of male and female emus in each group before and after a 7 day

treatment period.

Control female Starved female Control male Starved male

(ng/ml) (ng/ml) (ng/ml) (ng/ml)

(N=4) (N=5) (N=4) (N=3)

pre-treatment 0.80 ± 0.04 0.82 ± 0.06" 1.06 ±0.22 0.93 ± 0.07

treatment 1.45 ±0.36* 0.70±0.02*a 1.18 ±0.31 0.73 ±0.02

Values are means ± s.e.m. A n * indicates significant difference between groups (p = 0.008). The " indicates

significant difference over time (p = 0.0217).

The serum glucagon concentration was higher in the starved female group at the

completion of the treatment period (p = 0.036). The serum glucagon concentration did not

differ between the groups for the females pre-treatment, or the males pre-treatment or at

the completion of treatment. Over time there was no difference in glucagon within either

group for both sexes (Table 7.6).

Table 7.6: Serum glucagon concentrations of male and female emus in each group before and after a 7 day

treatment period.

pre-treatment

treatment

Control female

(pg/ml)

(N=3)

60.4 ±18.8

48.1 ±26.8*

Starved female

(pg/ml)

(N=5)

75.2 ±16.1

112.5 ±15.9*

Control male

(pg/ml)

(N=3)

46.1 ± 18.6

53.7 ± 29.8

Starved male

(pg/ml)

(N=3)

118.8 ±53.4

139.1 ±23.6



The insulin/glucagon ratio was lower in both the male and female starved groups at the

completion of treatment (p = 0.050 and p = 0.036 respectively). Over time there was a

tendency for the ratio to decrease in the male and female starved groups, however this was

not significant (p = 0.055 and p = 0.069 respectively). The control groups showed no

change in their insulin/glucagon ratios over time (Table 7.7).

Table 7.7: The insulin/glucagon ratio of male and female emus in each group before and after a 7 day

treatment period.


(N=3) (N=5) (N=3) (N=3)

pre-treatment 17.5 ±4.8 13.6 ±3.7 41.0 ±19.9 13.7 ±7.6

treatment 83.3 ±52.0* 7.0 ±1.4* 34.3 ± 10.9° 5.5±0.8a

Values are means ± s.e.m. A n * indicates significant difference between the female groups (p < 0.05). The a

indicates significant difference between the male groups (p < 0.05).

The serum thyroxine concentration was higher in the starved females at the completion of

treatment (p = 0.032). The concentration did not differ between the female groups at the

commencement of the pre-treatment period. For the male groups the thyroxine

concentration did not differ at the commencement of the pre-treatment period or the

completion of treatment. Over time there was no difference in thyroxine within either

group for both sexes (Table 7.8).

Table 7.8: Serum thyroxine concentrations of male and female emus in each group before and after a 7 day

treatment period.

pre-treatment

treatment

Control female

(nM/L)

(N=4)

2.59 ± 0.24

2.88 ±0.25*

Starved female

(nM/L)

(N=5)

3.53 ± 0.32

4.17 ±0.48*

Control male

(nM/L)

(N=4)

3.61 ±0.49

2.73 ± 0.79

Starved male

(nM/L)

(N=3)

4.37 ±0.29

4.32 ± 0.40



The serum triiodothyronine concentration was lower in the starved males than in the

control males at the completion of treatment (p = 0.028). The triiodothyronine

concentration did not differ between the female groups at either time-point or the male

groups pre-treatment. Over time the triiodothyronine concentration increased in the male

control group (p = 0.034). Over time the triiodothyronine concentration did not change for

the starved males or either female group (Table. 7.9).

Table 7.9: Serum triiodothyronine concentrations of male and female emus in each group before and after a 7 day treatment period.


(ng/mL) (ng/mL) (ng/mL) (ng/mL)

(N=4) (N=5) (N=4) (N=3)

pre-treatment 0.075 ±0.018 0.136 ±0.053 0.183 ±0.036" 0.120 ±0.061

treatment 0.090 ±0.033 0.092 ±0.019 0.223 ± 0.029*" 0.067 ±0.015*

Values are means ± s.e.m. A n * indicates significant difference between groups (p = 0.028). The " indicates significant difference over time (p = 0.034).

The triiodothyronine/thyroxine (T3/T4) ratio was lower in the starved than in the control

males at the completion of the treatment (p = 0.028). The T3/T4 ratio did not differ

between the female groups at either time-point, or the male groups pre-treatment. Within

groups there was no difference in the T3/T4 ratio for either sex over time (Table 7.10).

Table 7.10: Serum T3/T4 ratio (1 x 10"5) of male and female emus in each group before and after a 7 day

treatment period.

pre-treatment

treatment

Control female

(N=4)

7.26 ± 2.22

15.53 ±6.01

Starved female

(N=5)

3.59 ±1.79

2.09 ± 0.59

Control male

(N=4)

3.76 ±0.81

4.37 ±1.77*

Starved male

(N=3)

5.52 ± 2.39

2.96 ± 0.62*



The serum corticosterone concentration did not differ between the groups for either sex at

the commencement of the pre-treatment period or at the completion of treatment. Over

time there was no change in the concentration of corticosterone within any group (Table

7.11).

Table 7.11: Serum corticosterone concentrations of male and female emus in each group in each group before and after a 7 day treatment period.

pre-treatment

treatment

Control female

(ng/mL)

(N=4)

1.26 ±0.14

2.60 ±0.86

Starved female

(ng/mL)

(N=5)

1.84 ±0.88

1.41 ±0.15

Control male

(ng/mL)

(N=4)

0.82 ±0.12

1.47 ±0.19

Starved male

(ng/mL)

(N=3)

0.78 ±0.19

1.46 ±0.46


The serum B-hydroxybutyrate concentration did not differ between the groups for either

sex at the commencement of the pre-treatment period. At the completion of treatment the

B-hydroxybutyrate concentration was higher in the starved males than the control males,

but did not differ between groups for the females. Over time there was no change in the B-

hydroxybutyrate concentration within any group, however there was a trend for increased

B-hydroxybutyrate in the starved males (p = 0.055) (Table 7.12).

Table 7.12: Serum P-hydroxybutyrate concentrations of male and female emus in each group in each group

before and after a 7 day treatment period.


(mmol/L) (mmol/L) (mmol/L) (mmol/L)

(N=4) (N=5) (N=4) (N=3)

pre-treatment 0.62 ±0.26 0.34 ±0.17 0.05 ±0.03 0.03 ±0.02

treatment 0.32 ±0.31 0.83 ±0.40 0.29 ±0.23* 1.41 ±0.15*

Values are means ± s.e.m. A n * indicates significant difference between groups (p = 0.055)


The serum urea concentration did not differ between the groups for either sex pre-

treatment or for the females at the completion of treatment. At the completion of treatment

the urea concentration was lower in the starved males than the control males (p = 0.022).

Over time the urea concentration decreased within the starved female group and exhibited

a decreasing trend in the starved males (p = 0.022 and p = 0.055 respectively). Within the

control male and females groups there was no change over time in the concentration of

urea (Table 7.13).

Table 7.13: Serum urea concentrations of male and female emus in each group in each group before and

after a 7 day treatment period.


(mmol/L) (mmol/L) (mmol/L) (mmol/L)

(N=4) (N=5) (N=4) (N=3)

pre-treatment 1.70 ±0.30 1.38 ±0.27" 1.63 ±0.34 2.03 ±0.19

treatment 1.75 ±0.22 0.70 ±0.13" 1.78 ±0.26* 0.87 ±0.19*

Values are means ± s.e.m. A n * indicates significant difference between groups (p = 0.028). The " indicates

significant difference over time (p = 0.022).


4. Discussion

The hypothesis that N P Y m R N A expression would be elevated by short-term starvation in

male emus was not supported by the results. However, the hypothesis that VIP m R N A

expression would be unchanged was supported by the results. The hypothesis that short-

term starvation induces metabolic changes was partially supported by the results with one

week of starvation decreasing insulin concentrations in the female starved group below

that of both their pre-treatment concentration and the completion of treatment

concentration of the controls. In the females the glucagon concentration was also higher at

the completion of treatment in the starved group. This indicates that females may be more

sensitive to the effects of starvation than males. Effects on the thyroid hormones varied

between the sexes. The thyroxine concentration was higher in starved females at the

completion of treatment, while the triiodothyronine concentration and the T3/T4 ratio were

lower in starved males at the completion of treatment. This m a y again indicate a greater

sensitivity to starvation in the female e m u as the differences in the male values can largely

be explained by an increase in the triiodothyronine concentration of the controls at the

completion of the treatment period. Given the anorexia male emus exhibit when

incubating eggs, and the fact that females during the breeding season only exhibit a

reduced appetite, a greater resistance to starvation in male emus is a reasonable finding.

The corticosterone concentration did not increase in response to starvation in either sex,

indicating that seven days of starvation does not impose a large stress, as measured by

corticosterone concentration, on the emu.

The concentrations of metabolites were changed by starvation, and as with the hormone

concentrations these changes differed between the sexes. The concentration of (3-

hydroxybutyrate was higher in the starved male group at the completion of treatment.

Rather than indicating greater sensitivity to starvation, this may indicate an adaptation in

males that enables them to switch more quickly to the utilisation of adipose stores in

response to starvation than females. The starved female group showed a non-significant

increase in the concentration of p-hydroxybutyrate, and along with the initiation of

metabolic changes to conserve energy this m a y indicate that they employ a different

strategy to males in response to starvation. The concentration of urea decreased over time

in the female starved group and was lower at the completion of treatment in the male

starved group. This result was unexpected but can be explained in terms of the diet and

age of these animals. The animals used in this experiment were mature and received a


commercial diet containing 1 6 % protein. The reduction in urea concentration indicates

that the protein content of this diet exceeds the mature emus requirements. In the control

group the higher urea concentration is due to the need to breakdown and excrete excess

dietary protein.

The absence of changes in NPY mRNA expression in the mediobasal

hypothalamus/preoptic area or corticosterone concentration in response to starvation, and

the variable effects of starvation on metabolism implies two things. First, that the e m u is

more resistant to starvation than other species, and any effects of starvation on N P Y

expression or the concentration of corticosterone may only be observed after prolonged

periods. One week of starvation did not decrease either the subcutaneous or visceral

adipose reserves, although it did significantly decrease the liver weight as a proportion of

live-weight. The decreasing liver weight (more than 3 0 % lower) represents the utilisation

of stored glycogen reserves. A suitable period to examine the role of N P Y in the control of

appetite in the emu would have been such that the circulating concentration of

corticosterone was increased, the stored glycogen reserves were depleted by more than

3 0 % and catabolism of stored triglycerides from adipose tissue was initiated. Second, that

N P Y does not contribute significantly to the control of appetite in emus. Research in mice

has shown that multiple pathways are involved in the regulation of appetite and body

weight, and in the absence of N P Y , compensate to maintain adipose reserves in the correct

range (Erickson et al, 1996). It is therefore plausible that in the e m u one of these other

pathways is of greater importance to the control of appetite and bodyweight than N P Y .

Central pathways that could be of importance to the control of appetite in the e m u include

those involving cocaine and amphetamine-regulated transcript (CART), orexin/hypocretin,

corticotrophin-releasing hormone (CRH), proopiomelanocortin ( P O M C ) and agouti-related

protein (AGRP).

In conclusion, subjecting emus to one week of starvation does not increase NPY gene

expression or induce the range of hormonal changes that are associated with starvation in

other species. This m a y be due to a greater tolerance to starvation in emus. Given that

male emus exhibit anorexia for a period of eight weeks during incubation, it is not

surprising that a single week of starvation failed to induce the classical starvation

responses. The absence of any change in N P Y m R N A expression in response to short-

term starvation could also be due to a more limited role of N P Y in appetite control than is


observed in other species. This experiment did not clarify the roles of N P Y and VIP in

appetite control or the possibility of mediation of NPY's effects on appetite by

corticosterone. Performing this experiment over a longer time period may have yielded

more information on the control of appetite in the emu, as would comparisons between the

expression of N P Y during the breeding season (minimal appetite) and the summer months

(maximal appetite).

162

General Discussion


The e m u has great potential as a commercial livestock species. The major drawback to

achieving this potential is the seasonal decrease in appetite and adiposity that they exhibit

in response to decreasing photoperiod. Decreased appetite, and particularly the ensuing

decreased adiposity, are a problem because fat is the most profitable component of the

carcass. In a commercial production scenario the decreased appetite and adiposity,

restricts the viable window for slaughter to a brief period during the summer months. As a

consequence of this restriction the availability of e m u products (i.e. meat, leather and fat)

are limited and market development is hampered. The aim of this thesis has been to

improve the understanding of h o w appetite and adiposity are controlled in the emu.

Improving our understanding of these controls is essential for identifying mechanisms to

manipulate appetite and adiposity, and for developing techniques to assess the

susceptibility of the e m u to these manipulations.

Emus might have different mechanisms to control appetite and adiposity than those

decribed in other species. During the course of this thesis emus have demonstrated a

striking resistance to the manipulation of both appetite and adiposity (see Chapters 4 and

5) and tolerance to short-term starvation (see Chapter 7). E m u s are also well adapted to

survive prolonged negative energy balance during the anorexia of incubation (see Chapter

6). E m u s have evolved in a harsh environment in which is food is often scarce (Davies,

1978). As such, the unique features of the emu demonstrated during the course of this

thesis m a y result from adaptations to this harsh environment. To cope with food scarcity

and the demands of reproduction the emu has evolved efficient mechanisms to maintain

energy reserves at an appropriate level. These mechanisms include strategies for coping

with starvation that diminish the stress response and appetite stimulation, and thereby

minimise discomfort and the negative effects of continuously elevated stress hormones on

the animal (see Chapter 7). The efficient and rigid control of appetite and adiposity allows

the e m u to resist treatments that are known to alter appetite and adiposity in other species,

such as, dexamethasone treatment (see Chapter 5) and the consumption of high fat diets

(see Chapter 4). As such, it may not be possible to maintain high appetite and adiposity in

the e m u during the breeding season.

My work has focused on how to prevent appetite and adiposity decreasing. It is possible

that this is the wrong approach for tackling the problem of a limited slaughter window for

producers. I assumed that appetite was decreasing during the reproductive period and that


the emus could not meet their requirements during this period when compared to the large

amount of food ingested over summer. In fact, the increase in appetite during summer may

be part of a strategy of over feeding to increase energy reserves and support the demands

of reproduction during winter. Premigratory fattening is an analogous situation where

appetite is increased to increase adipose reserves and enable migrating species to cover

large distances when a food supply is unreliable or unavailable (Stevens, 1996). For a

species to survive it must reproduce (Jones, 1999). Therefore factors that influence the

success of reproduction, such as the control of energy reserves, come under intense

selection pressure. Adaptations that have evolved under such intense selection are unlikely

to be overcome with ease. Therefore, it may be better to investigate the mechanism

controlling the stimulation of appetite at the completion of incubation. Studies into the

rapid and large increase in food intake of the male emu observed at the cessation of

incubation would be valuable and could provide feasible mechanisms for increasing

appetite earlier and thereby extending the slaughter window.

Natural selection has encouraged the evolution of photoperiod-related controls of appetite

and adiposity in the emu that strongly resist interference (Blache & Martin, 1999; Blache

et al, 2001b). The photoperiodic control of appetite most likely involves a complicated

network involving interactions between hypothalamic neuropeptides, peripheral metabolic

and reproductive hormones, and photoperiod-related factors. The photoperiodic control of

appetite during the reproductive period will be difficult to characterise fully and m a y be

impossible to override. A promising approach to manipulate appetite and adiposity during

the reproductive period would be to alter the perception of photoperiodic changes by the

emu. In mammals, melatonin is a key component of photoperiodicity and its

administration can inhibit reproduction (Edmonds & Stetson, 1994; Lincoln, 1994; Horton

& Yellon, 2001). However, in birds, it appears that melatonin may not be involved in the

control of reproduction by photoperiod, although, the role of melatonin in emus has never

been investigated (Meyer, 1998). Other factors that could provide insights into the

photoperiodic control of appetite and potential mechanisms for disrupting it include;

norepinephrine, gonadotrophin-releasing hormone, and proopiomelanocortin ( P O M C ) and

agouti-related protein (AGRP) (Kriegsfeld et al, 2000; Mercer et al, 2000a & 2000b;

Imundo et al, 2001), further investigations of these pathways could lead to ways to

manipulate the effect of photoperiod on appetite in emus.


In conclusion, I have explored several avenues to prevent the loss of appetite and adiposity

during the breeding season. Methods that are known to increase appetite and adiposity in

other species have little or no effect in emus. It is possible that investigations into the

mechanisms that trigger the rapid and large increase in appetite and adiposity after the

breeding season could yield new approaches for manipulating appetite and adiposity in the

e m u to extend the slaughter window. Another area for investigation that could provide

methods for manipulating appetite and adiposity is the neural basis for the control of

appetite by photoperiod.

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Appendix 1 - Abbreviations 202

Abbreviations used in the thesis

ACTH

AGRP

amp

ANSA

ATP

Bo

BMI

BSA

CART

cDNA

CER

CH3COONa

C8HiiN2Na03

CP

cpm

CRH

CSIRO

CTP

db/db

DNA

EDTA

GPB

GTP

IU

KI

LepR

LiCl

MBH

mRNA

NaCl

Adrenocorticotropin

Agouti-related protein

Ampicillin

8-anilino-l -naphthalene sulphonic acid

Adenosine triphosphate

Zero binding

Body mass index

Bovine serum albumin

Cocaine and amphetamine-related transcript

Complementary D N A

Cerebellum

Sodium acetate

Barbital sodium

Choroid plexus

Counts per minute

Corticotrophin-releasing hormone

Commonwealth Scientific and Industrial Research Organisation

Cytocine triphosphate

Diabetes genotype

Deoxyribonucleic acid

Ethylenediaminetetra acetic acid

Gelatin phosphate buffer

Guanidine triphosphate

International units

Potassium iodide

Leptin receptor

Lithium chloride

Mediobasal hypothalamus

Messenger ribonucleic acid

Sodium chloride

Appendix 1 - Abbreviations

NaN3

NaOH

Na2HP04

NaH2P04.2H20

Na2S205

NGPS

NH4OAC

NIH

NPY

NRS

NSB

ob/ob

PBS

PCR

PEG

PIPES

POA

POMC

PS

RACE

RIA

RERDC

RPA

rpm

RT-PCR

SDS

s.e.m.

SSC

Taq

TC

T E M E D

T R H

Sodium azide

Sodium hydroxide

Disodium hydrogen orthophosphate

Sodium dihydrogen orthophosphate

Sodium metabisulphite

Normal guinea pig serum

Ammonium acetate

National Institute of Health

Neuropeptide Y

Normal rabbit serum

Non-specific binding

Obese genotype

Phosphate buffered saline

Polymerase chain reaction

Polyethylene glycol

Piperazine-N,N'-bis[2-ethane sulfonic acid]

Preoptic area

Proopiomelanocortin

Pituitary stalk

Rapid amplification of c D N A ends

Radioimmunoassay

Rural Industries Research and Development Corporation

Ribonuclease protection assay

Revolutions per minute

Reverse transcription polymerase chain reaction

Sodium dodecyl sulfate

Standard error of the mean

Sodium chloride sodium citrate

Taq polymerase enzyme

Total counts

N,N,N' ,N' -tetra-methyl-ethylenediamine

Thyroid releasing hormone

Appendix 1 - Abbreviations

U T P Uracil triphosphate

U W A University of Western Australia

VIP Vasoactive intestinal peptide

V L D L Very low-density lipoprotein

V M H Ventromedial hypothalamic nucleus

Appendix 2 - List of suppliers and feed composition 205

Alphabetical list of suppliers

Consumables, chemicals and equipment

Ambion, Inc. 2130 Woodward Austin TX 787744-1832 USA http://www.ambion.com/ Distributed in Australia by Gene Works Pty Ltd

Amersham Biosciences Pty Ltd Unit 1, number 22 Hudson Avenue Castle Hill N S W 2154 AUSTRALIA

Beckman (now Beckman Coulter Australia Pty Ltd) Unit C, 24 College Street Gladesville NSW 2111 AUSTRALIA http ://w w w .beckman .com/

Biogenesis Immuno Diagnostics (now Biogenesis Ltd) Technology Road Poole BH17 7DA UK http://www.biogenesis.co.uk/

BioRad Laboratories Pty Ltd Unit 1, Block Y Regents Park Industrial Estate 391 Park Road Regents Park N S W 2143 AUSTRALIA http://www.bio-rad.com/

Boehringer Mannheim (acquired by Roche Products Pty Ltd) 4-10 Inman Road Dee Why N S W 2099 AUSTRALIA http://www.roche.com/

Clontech (now B D Biosciences Clontech) 4 Research Park Drive Macquarie University Research Park North Ryde N S W 2113 AUSTRALIA http://www.clontech.com/

Endocrine Sciences Products (now Endocrine Sciences RIA Reagents) 4301 Lost Hills Rd Calabasas C A 91301 USA

Genesearch Pty Ltd 14 Technology Drive Arundel Q L D 4214 AUSTRALIA

GeneWorks Pty Ltd 39 Winwood Street Thebarton S A 5031 AUSTRALIA

http://www.ambion.com/

http://www.biogenesis.co.uk/

http://www.bio-rad.com/

http://www.roche.com/

http://www.clontech.com/


GIBCO (now Invitrogen Life Technologies) In Israel distributed by R H E N I U M Ltd PO Box 3580 91035 Jerusalem ISRAEL http://www2.lifetech.com/

Hybaid (now Thermo Hybaid) Thermo Biosciences G m b H Sedanstr. 18 D-89077 Ulm GERMANY http ://ww w.hybaid- gmbh.com/

Ilium Veterinary Products (supplied by Troy Laboratories Pty Ltd) http ://w w w .troylab .com, au/

Integrated Sciences PO Box 196 East Kew Melbourne VIC 3102 AUSTRALIA http ://www .integratedsci.com. au/

Linco Research, Inc. 14 Research Drive St Charles Missouri 63304 U S A

Medisense (produced by Abbott Laboratories) Abbott Diagnostics Division

PO Box 394 North Ryde N S W 2113 AUSTRALIA http://www.medisense.com/ http://www.abbottlabs.com.au/

N E N Life Science 549 Albany Street Boston M A 02118-2512 USA

New England Biolabs http ://w w w .neb .com/ Distributed in Australia by Genesearch Pty Ltd

Packard (now PerkinElmer Life Sciences) PerkinElmer Pty Ltd 16-18 Kingsley Close Rowville Melbourne VIC 3178 AUSTRALIA

Pharmacia Biotech (now Amersham Biosciences Pty Ltd)

Promega Corp. Australia 37 Nelson Street Annandale N S W 2038 AUSTRALIA http ://www.promega.com/

QIAGEN Pty Ltd PO Box 25 Clifton Hill VIC 3068 AUSTRALIA http://www.qiagen.com/

Sarstedt Australia Pty Ltd PO Box 90 Ingle Farm SA 5098 AUSTRALIA http://www.sarstedt.com/

http://www2.lifetech.com/

http://gmbh.com/

http://integratedsci.com

http://www.medisense.com/

http://www.abbottlabs.com.au/

http://www.promega.com/

http://www.qiagen.com/

http://www.sarstedt.com/

Appendix 2 - List of suppliers and feed composition

Sigma (now Sigma-Aldrich Pty Ltd) Sydney AUSTRALIA http://www.sigma.aldrich.com/

Stratagene 11011 N. Torrey Pines Road La Jolla C A 92037 U S A http://www.stratagene.com/ Distributed in Australia by Integrated Sciences

Tel-test Inc. PO Box 1421 Friendswood T X 77546 U S A http://www.isotexdiagnostics.com/ Distributed in Australia by Gene Works Pty Ltd

Turner Design Inc 845 W. Maude Avenue Sunnyvale C A 94085 USA http://www.turnerdesigns.com/

Virbac 15 Pritchard Place Peakhurst N S W 2210 AUSTRALIA http://www.virbac.com.au/

Whatman International Ltd Whatman House St Leonards Road 20/20 Maidstone Kent ME16 0LS U K http://www.whatman.com/

Diet components and commercial feeds

Bayer PO Box 1045 3 De Laeter Way Bentley WA AUSTRALIA http://www.baver.com.au/

Davison Industries Pinjarra WA AUSTRALIA

Education Colours Pty Ltd PO Box 657 Croydon VIC 3136 AUSTRALIA

Glen Forrest Stockfeeders 3150 Great Eastern Highway Glen Forrest WA6076 AUSTRALIA

http://www.sigma.aldrich.com/

http://www.stratagene.com/

http://www.isotexdiagnostics.com/

http://www.turnerdesigns.com/

http://www.virbac.com.au/

http://www.whatman.com/

http://www.baver.com.au/


Feed composition

Glen Forrest Stockfeeders emu breeder pellet:

1 5 % protein

7.6% crude fibre

3.7% fat

providing 10.5MJ/kg of metabolisable energy.

209

Appendix 3

Response to comments made by thesis examiners

Page

Chapter 2 210

Chapter 4 212

Chapter 5 214

Chapter 6 218

Chapter 7 222

Appendix 3 - response to comments made by examiners 210

Chapter 2.

Question 1. Were feed refusals sub-sampled and analysed to determine actual intake i.e. was

there any scope for birds to select differently from diet specifications?

Refusal margins were not calculated as the amount fed to each bird was adjusted continuously

in response to changes in intake. Refusals were not sub-sampled as the diet was pelleted and

the components had been finely ground making differential selection less likely. The two

diets created for experiment 1 (Chapter 4) were also sifted at each weighing and refilling to

remove any powder that had accumulated, further reducing the opportunity for differential

selection of the diet components.

Question 2. Corticosterone assay - why were different sized tubes used for samples and

standards?

As the standards were not extracted they were set up in a tube appropriate to the volume used

for the assay. The extraction of corticosterone was performed in larger tubes that were

appropriate for the volume of iso-octane/hexane used the resulting organic layer was then

transferred to the smaller tubes for drying down and continuation of the assay.

Question 3. Glucagon assay - what is aprotinin? why was normal guinea pig serum (NGPS)

used?

Aprotinin is added to the assay to protect against protease activity. N G P S was added to the

assay to aid precipitation.

Question 4. Insulin assay - why was 0.1% gelatine added to the assay buffer?

Insulin may be present in B S A preparations, therefore gelatine was added in preference to

B S A to prevent confounding of the results by insulin contamination. Additionally the use of

gelatine in the assay buffer was recommended in the original assay method.

Question 5. LH assay - why were fractions eluted into buffer plus BSA?

The B S A stops the radioactively labelled L H from sticking to the tube.


Question 6. T3/T4 assays - why was stripped plasma added to the standards in both assays?

why were glass tubes used?

Stripped plasma was added to the tubes to provide a blank effect for plasma in the assay.

Glass tubes were used because the assay was performed in a small volume and with glass

tubes droplets don't stick to the sides and therefore give less error.

Question 7. Why wasn't glucagon assay 2 with 66% binding repeated?

The components used in the glucagon assay were mainly purchased from a commercial

supplier and the cost of repeating this assay was prohibitive.


Chapter 4.

Question 1. Figure 1.1 suggests through spring and summer that males gain significant live

weight, why then did neither group gain weight during the experiment?

The birds used in this experiment were caught fortnightly for weighing and bleeding. The

handling involved in this process, in particular the weighing procedure, can be stressful for the

bird. This stress may have had a negative effect on appetite in these birds. Indeed, a decrease

in food intake was observed in all birds after handling. Their food intake slowly recovered to

pre-handling levels over two weeks, at which time they were handled again. In subsequent

experiments weighing of birds was kept to a minimum to try and avoid this effect.

Question 2. As neither group gained weight changes in total fat would have to occur through

a decrease in the weight of another tissue, is this suggested anywhere in the literature? Or

were you never going to get a response because weight did not change?

The literature does not indicate that adipose tissue weight can increase when weight is

constant. However, there is evidence that the type of fat consumed can influence the oxidation

rate of fat in chickens and therefore the partitioning of energy into lean or fat tissue (Crespo &

Esteve-Garcia, 2002; N e w m a n et al, 2002). Saturated fats induce greater abdominal adipose

deposition than unsaturated fats in chickens (Vila & Esteve-Garcia, 1996; Sanz et al, 1999 &

2000). As such, a diet high in saturated fat could cause adiposity to increase despite only

moderate changes in weight. In combination though the absence of increases in weight gain in

this experiment and the use of unsaturated fat in the diet did make it unlikely that an increase

in adiposity would be observed.

Question 3. What are the hormonal impacts on synthesis, degradation and accretion rates in

fat and muscle?

The key hormones involved in the storage and mobilisation of energy reserves are insulin,

glucagon and glucocorticoids. They affect synthesis, degradation and accretion rates of fat

and muscle through various mechanisms.

1) Insulin:

- in the liver increases glucose uptake, inhibits gluconeogenesis, and increases

lipogenesis and the release of V L D L into the peripheral circulation (Stevens, 1996),

- in adipose tissue, increases glucose uptake, inhibits free fatty acid release, stimulates


the conversion of glucose to fat and stimulates lipoprotein lipase concentrations

(Griminger, 1986; Hazelwood, 1986), sparing the triglyceride stores and increasing

adipose deposition,

- in muscle increases the rate of glucose uptake, facilitates the formation of glycogen

from glucose, inhibits glycogen mobilisation and stimulates the uptake of amino acids

(Bentley, 1998).

2) Glucagon:

in the liver mobilises energy reserves by inhibiting lipogenesis and stimulating

gluconeogenesis (Stevens, 1996),

- in adipose tissue stimulates lipolysis (Stevens, 1996), releasing free fatty acids into the

peripheral circulation (Braganza et al, 1973),

- in muscle tissue mobilises glycogen and amino acids (Bentley, 1998).

3) Glucocorticoids:

- in the liver stimulate lipogenesis and gluconeogenesis (Griminger, 1986; Stevens,

1996),

- in adipose tissue stimulate lipolysis (Griminger, 1986; Stevens, 1996),

- in muscle mobilise protein and deaminate amino acids that are then used as a substrate

for gluconeogenesis in the liver (Bentley, 1998).

Question 4. Does glucagon have lipolytic potential in emus as in other avian species?

Glucagon was elevated in the high fat group, therefore was intake depressed because of the

high fat diet and glucagon increasing lipolysis and therefore the circulating energy levels

were very high and intake was suppressed?

It has not been assessed in the e m u but it is likely that glucagon does have the same lipolytic

potential observed in other avian species. The results for glucagon indicate a trend for

elevated glucagon in the high fat group at a single time point. This is at the end of the diet

changeover period when the birds showed the largest reductions in food intake. As such, the

raised glucagon levels are more likely to be a consequence of decreased food intake, rather

than a cause. It is possible that the decreased feed intake led to an increase in glucagon to

stimulate lipolysis and meet the energetic requirements of the animals.


Chapter 5.

Question 1. How do terminal fat scores relate to actual measurements of carcass adiposity?

Furthermore a correlation analysis would help to calibrate the subjective scoring system for

future use.

The fat scoring system used has been previously shown to correlate well with adiposity of emu

carcasses as assessed by ultrasound (Mincham et al, 1998). The results for fat scoring

performed during experiment 2 (Chapter 5) exhibit a strong positive relationship between fat

score and the percentage total adiposity (correlation coefficient = 0.918) (Figure A3.1).

Analysis of variance indicates that this relationship is significant (F ratio = 53.48, => p value

< 0.001).

10 15

Total adiposity (%)

20

Figure A3.1: Correlation analysis of the relationship between fat score and percentage total adiposity of female in experiment 2 (saline treated = open circles and dexamethasone treated = solid circles).

emus


Question 2. How representative are single LH determinations? What are the LH secretory

dynamics in female emus? Why not measure ovarian steroids? And/or FSH?

In chickens, adult cockerels but not laying hens have been shown to have episodic L H release

(Wilson & Sharp, 1975). In turkeys L H secretion by females differs between photosensitive

and photostimulated birds. Photosensitive birds have L H pulses of low frequency and high

amplitude with low baseline L H concentrations and photostimulated birds have a high baseline

L H concentration with few pulses of low amplitude and short duration (Chapman et al, 1994).

In emus serial blood sampling did not show any evidence of L H pulsatility in the female emu

(Van Cleeff, 2001). As such, single determinations can be considered to accurately represent

L H concentration in the emu. Ovarian steroids and F S H were not measured due to the cost

and time that would have been required to develop and validate the assays for emus.

Question 3. Is there a reference for reduced feed intake leading to reduced ovarian weight in

emus/birds (over time course here of 3 weeks with appetite reduced by 30-70%)?

Feeding chickens a diet high in zinc or fasting them for 10 days decreases food intake and

ovary weight (McCormick & Cunningham, 1987). Restricted feeding of broilers to 4 5 % of ad

libitum also results in lower ovary weights (Waddington & Hocking, 1993). However,

glucocorticoid administration has been shown to decrease ovary weight in the hen in the

absence of changes in live weight or food intake (Williams et al, 1985). As such, the

suppression of ovary development observed in experiment 2 (Chapter 5) may be due initially

to treatment with dexamethasone and compounded by the reduction in food intake observed at

the end of the experiment.

Question 4. Lipid infiltration of the emu liver in response to dexamethasone treatment -

discuss/reference more fully.

Dexamethasone treatment can cause lipid infiltration of the liver (Franco-Colin et al, 2000;

Letteron et al, 1997). Dexamethasone causes lipid infiltration in several ways:

1) increasing the activity of phosphatidate phosphohydrolase and thereby the synthesis of

neutral lipids from glycerol phosphate (Pittner et al, 1995),

2) inhibiting medium- and short-chain acyl C o A dehydrogenation and hepatic lipid

secretion (Letteron et al, 1997)


1) inducing haptoglobin production, an acute phase protein associated with hepatic

lipodosis (Higuchi etal, 199A).

The increased liver weight observed following chronic treatment with dexamethasone during

experiment 2 (Table 5.2) and the associated difference in appearance of the liver between the

two groups (Figure A3.2) are consistent with lipid infiltration.

Figure A3.2: Liver tissue from female emus following chronic treatment with either saline (left) or dexamethasone (right).

Question 5. Increased blood glucose result is not discussed. Treatment reduced the

insulin:glucose ratio? Why did this not effect NPY?

The increased blood glucose resulted from the action of dexamethasone to inhibit peripheral

oxidation and utilisation of glucose. The elevation of blood glucose, in the absence of changes

in the concentration of insulin, did result in a decrease in the insulimglucose ratio. Elevated

insulin concentrations act to decrease N P Y gene expression (Schwartz et al, 1991 & 1992).

Therefore it is unlikely that insulin was influencing N P Y expression in this experiment. In the

arcuate nucleus glucose-sensitive neurons overlap with NPY-containing neurons (Muroya et

al, 1999). A decrease in central glucose concentration activates these neurons causing them to

release N P Y and thereby stimulate feeding and normalisation of peripheral and central glucose

concentrations. A s peripheral glucose concentrations were increased in experiment 2, it is

unlikely that they would have stimulated N P Y by this mechanism. Acute stimulation of blood

glucose concentration is associated with elevations of N P Y gene expression and corticosterone

concentration (Wang et al, 1999). As corticosterone was depressed by dexamethasone


treatment in experiment 2, it is possible that the absence of an effect of glucose on NPY

expression reflects a requirement for corticosterone to potentiate any effects of elevated

glucose on N P Y gene expression.

Question 6. Why did animals fed ad libitum not gain weight?

This experiment involved weekly handling of the animals for injections and bleeding, and as

for experiment 1 this regular handling was most likely a significant stressor for the birds that

may have reduced their food intake. The pattern of reduced feed intake after handling

observed in experiment 1 was not seen during this experiment. However, the greater

frequency of handling may have prevented appetite recovering in the interim period.


Chapter 6.

Question 1. Compare food intake of incubating and non-incubating birds to "control" birds

during the summer.

The food intake of non-incubating emus during autumn (short daylength) and summer (long

daylength) on-farm, and under equivalent photoperiod treatments, was higher in both

experiments in the summer/long day groups (Blache & Martin, 1999; O'Malley, 1996).

Question 2. Does blood sampling during incubation constitute a major disturbance? Do

emus abstain while sitting but get up at certain times to feed?

Blood sampling does not constitute a major disturbance during incubation. Birds remained on

the nest during the procedure and generally showed no response to the handling involved. The

birds abstain from eating while sitting and very rarely leave the nest. During the incubation

period they exhibit anorexia.

Question 3. Also the time of day of killing might be important? Could there be diurnal

changes in NPY expression controlling the moment-to moment drive to eat?

The birds were always killed in the morning. As such, diurnal changes in N P Y expression

should not have confounded the results of this experiment. Diurnal changes in N P Y

expression could control the moment-to-moment drive to eat in emus. Investigation of the

eating patterns of emus has revealed that they only eat during the light period and that the

difference in food intake observed between birds on L D and S D is due to differences in the

time spent eating rather than in the frequency of meals (Blache & Martin, 1999). O n LD, but

not SD, the final 4 hours of light was associated with a two-fold decrease in feeding

frequency. In rats, N P Y gene expression in the hypothalamus increases during the light phase,

decreases just prior to the onset of the dark phase and remains low throughout the dark phase

(Xu et al, 1999). Food intake in rats is highest during the dark phase and lowest during the

light phase (Lu et al, 2002). Therefore, increased N P Y expression during the light phase may

be a response to negative energy balance during this period that ensures appetite is high when

the dark phase begins. The decreased N P Y expression during the period of maximal appetite

can be explained by increases in peripheral signals such as insulin and glucose that result from

eating. If N P Y gene expression shows diurnal changes that regulate appetite in a similar


manner to rats it could be expected that N P Y gene expression would be highest during the

early hours of the morning and maintained at a lower constant level throughout the day.

Question 4. How was it possible to quantify lanes 1 and 2 ?

It is possible that these lanes are saturated, the data from the image analysis program did

however appear reasonable and the data was therefore retained to keep the birds in the

incubating group at a reasonable number. Leaving the data for these lanes out of the Mann-

Whitney U test does not change the interpretation of the results (Table A3.1).

Table A3.1: The m R N A expression of NPY and VIP relative to p-actin of incubating (n=3) and non-incubating (n=6) male emus.

m R N A Incubators Non-incubators

NPY/p-actin 0.68 ± 0.075 0.53 ±0.083

VIP/p-actin 0.66 ± 0.030* 0.47 ± 0.055*

Values are means ± s.e.m. An * indicates significant difference between groups (p = 0.019).

Question 5. The trend to increased NPY could relate to the increased corticosterone,

prolactin or reduced insulin:glucagon ratios?

Using correlation analysis to investigate relationships between N P Y gene expression and

either corticosterone, prolactin or the insulimglucagon ratio, it was determined that strong

relationships exist between N P Y and both corticosterone and prolactin, but not the

insulin:glucagon ratio. There is a strong positive relationship between corticosterone and

N P Y gene expression in the non-incubating group (Table A3.2). Analysis of variance

indicates that this relationship is significant. The data for the incubating group is confounded

by the small number of animals used, it appears though that four out of the five data points

correspond closely to the relationship observed in the non-incubating group (Figure A3.3).

For prolactin the results indicate a weak positive relationship with N P Y expression in the non-

incubating group (Table A3.2) and a strong negative relationship in the incubating group.

Analysis of variance indicates that this relationship is significant. This raises the interesting

possibility that prolactin could have a role in regulating NPY's effects on appetite that is

dependent upon the reproductive status of the animal. The insulin:glucagon ratio showed no

evidence of a relationship with N P Y gene expression for either group.

Appendix 3 - response to comments m a d e by examiners 220

Table A3.2: Correlation analysis comparing the expression of N P Y relative to p-actin in incubating and non-incubating male emus with corticosterone and prolactin concentrations and the insulin:glucagon ratio.

Group and hormone interaction

Non-incubators -corticosterone (ng/ml)

Incubators -corticosterone (ng/ml)

Non-incubators -prolactin (ng/ml)

Incubators -prolactin (ng/ml)

Non-incubators -insulin: glucagon ratio

Incubators -insulin: glucagon ratio

Correlation coefficient

0.994

-0.692

0.538

-0.899

-0.542

0.469

F ratio

256.6

2.759

1.63

12.7

0.53

0.85

p value

< 0.001

>0.05

>0.05

<0.05

>0.05

>0.05

0.90

0.80 c o 8 0.70 & X C u

0.60-

•fj 0.50 g a • g 0.40

£ 0.30

.2 0.20

0.10 -

0.00

0.5

R2 = 0.987

1 1.5 2 2.5

Corticosterone concentration (ng/ml)

— i

3.5

Figure A3.3: Correlation analysis of the relationship between N P Y gene expression relative to P-actin and corticosterone concentration (non-incubators = open circles and incubators = solid circles).


Question 6. Would differences have been more significant with greater n values?

If more animals had been available to use in the experiment the differences observed between

the groups would have been more likely to be significant. Greater n values in each group

would have reduced the confounding effects of the high variability observed between birds for

many of the parameters measured.


Chapter 7.

Question 1. One week of starvation seems severe. Can this be justified? Did this and all

experiments receive ethical approval beforehand?

The application of one week of starvation is justified given the much more extreme voluntary

starvation that emus undergo during incubation. All experiments received approval from the

U W A animal ethics committee and were performed in accordance with the animal ethics and

welfare guidelines given by U W A . Chapter 2 has been amended to reflect this.

Question 2. Was the carcass analysis performed on the same subset of birds used for the

hypothalamic NPY measurement?

Yes, the material and methods section of chapter 7 has been amended to more clearly reflect

this.

Question 3. Another possibility worthy of discussion is that the RPA method of measuring

mRNA in blocks of hypothalamic tissue fails to detect small but biologically significant

changes in expression within specific nuclei, or in subpopulations of neurons within nuclei.

This is a fair criticism of the technique, for detecting genes that are weakly expressed or subtle

changes in gene expression it is not an appropriate method unless the region where the gene is

expressed is very accurately dissected. In Japanese quail fasted for 24 hours and chickens

fasted for 48 hours or chronically feed restricted R P A detected for quail 1.5 and for chicken 2

fold increases in N P Y gene expression (Boswell et al, 2002; Boswell et al, 1999; Boswell et

al, 1999b). As such, R P A is a suitable technique for studying N P Y expression in the emu

hypothalamus.

appetite and adiposity of the emu (dromaius novaehollandiae)

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