A Thyroid Hormone Based Therapy to Restore Brain ...

A Thyroid Hormone Based Therapy to

Restore Brain Maturation Following

Foetal Growth Restriction

A thesis submitted in fulfilment of the requirements for the degree of

Doctor of Philosophy

Delphi Eleni Kondos-Devcic

BSc (Hons) Monash University

School of Health and Biomedical Sciences

College of Science, Engineering and Health

RMIT University

September 2020

Declaration

I certify that except where due acknowledgement has been made, the work is that of the author alone;

the work has not been submitted previously, in whole or in part, to qualify for any other academic

award; the content of the thesis is the result of work which has been carried out since the official

commencement date of the approved research program; any editorial work, paid or unpaid, carried

out by a third party is acknowledged; and, ethics procedures and guidelines have been followed.

I acknowledge the support I have received for my research through the provision of an Australian

Government Research Training Program Scholarship.

Delphi Eleni Kondos-Devcic

05 September 2020

RMIT University

I

Dedicated to my parents

Stavroula Kondos & Dusan Devcic

II

“Above all, don’t fear difficult moments. The best comes from them.”

Rita Levi-Montalcini, Nobel Laureate.

III

Acknowledgements

My PhD has been a transformative journey and every step contained valuable lessons. The last four

years have shaped the woman I am today. This journey has been guided and supported by the

following individuals and I am eternally grateful for my experiences with them. First and foremost,

I thank my supervisors, Associate Professor Mary Tolcos, Professor David Walker and Associate

Professor Flora Wong. Without your combined expertise and encouragement I would not have

accomplished this amazing milestone in my life.

Mary, when I first met you I was an undergraduate who had never in my wildest dreams imagined I

was capable of completing a PhD. Thank you for encouraging me and believing in my abilities. It has

been one of the most difficult and at the same time fulfilling adventures of my life so far! You have

taught me to keep calm when things go wrong and that a good solution can ALWAYS be found for

any problem. Thank you for teaching me to pay attention to the details, as I am now a much better

writer and presenter because of this. I’m also very glad to have been your first RMIT student and will

carry many great memories of our time together at RMIT, including countless Vietnamese lunches

across the road!

David, thank you for shining a constant positive light during my PhD and boosting my confidence

that I was on the right track, even when I didn’t feel it myself. Your ‘big-picture’ approach has helped

me to think laterally and expand my mind, and this skill will no doubt serve me well throughout my

life. Thank you for your support of my role in the SOBR committee, where all of our events

were enhanced because you attended and were on our judging panels. Your feedback ahead of all my

presentations was very much appreciated, and I learnt many valuable presenting skills from you.

Flora, thank you for offering me your wealth of clinical expertise. You helped me to understand the

clinical implications of my research more fully. Your advice and input was very much appreciated.

Although we were at different institutions, you always made time to reply to my e-mails. I also very

much enjoyed attending the 3rd JENS conference in the Netherlands with you in 2019.

Dr. Angela Cumberland not only assisted me in setting up my animal model at RMIT, but was an

amazing support system and friend outside of the lab. Angie, thank you for teaching me to be

meticulous and resourceful in the lab, and for taking me on sanity-breaks across the road

to Lindt or Nevski café for toasties. You are the meaning of ‘girl power’!

IV

Ms Madhavi Khore Madi, thank you for all your help in setting up the lab when we first arrived at

RMIT. Your organisational skills, amazing work ethic, and helping hands with my animals made the

whole process easier for me and I am very grateful for that. Thank also for your friendship!

Dr. Azu (Aminath) Azhan Azu, thank you for being an important (sparkly/sequined) role model for

me during my PhD. I watched you complete your own PhD while also excelling in your personal life,

including being appointed president of the Rotary club of Melbourne Park. You have inspired me to

be a strong woman who makes change in the world.

Tolcos/ Walker Laboratory Group Thank you to lab members of the Tolcos/Walker group both

past and present for their support during my PhD: Courtney, James, Sebastian, Issy, Nhi, Bobbi,

Abdul, Emily, Ginevra, Ryan. It has been an absolute pleasure to be part of such a supportive and fun

team. Courtney, a special thank you for all of the runs/ gazelle sprints around Princes Park!

The RMIT Crew Simone, Alita, Bashira, Christian, Emma, Jono, Ivan, Kurt, Alec, Chris,

Paris, Hanife, Mingdi, Maurice, Sherouk, thank you for the chats, support and laughs during the last

4 years. A wonderful bunch of people who I am now lucky enough to call my friends!

The Hudson Crew Shreya, Kelsee, Lara, Nadia B. and Nadia H., A collective of strong independent

women who have left such a wonderful impression on my life and on my PhD experience. Shreya

and Kelsee, your constant support and kind words during challenging times will never be forgotten.

Panel Members Thank you to my panel members, who devoted their time and energy, and gave me

valuable advice throughout my candidature and final thesis submission: A/Prof. Samantha

Richardson and Dr.Luba Sominski, A/Prof Timothy Regnault, Dr Thomas Schmitz.

My animals I feel deep respect and gratitude to the rats who were sacrificed for this research. These

animals have allowed me to uncover valuable knowledge which will ultimately benefit our society.

It is very sad to sacrifice animals and I think about my rats often. Their essential contribution to

advances in novel therapies should never be forgotten.

Family and friends Thank you Paloma for being the best friend anyone could ask for. You celebrated

even my smallest achievements and provided constant support, while also being an ultimate stage-

mum at all of my candidature milestone presentations. Amelia, my sister from another mister, thank

you for all of the adventures and for picking me up with warmth when I had fallen down.

V

Lisa, Shalini, Liv, Audrey, Jess, Molly, Ally & Sarah, your unwavering support, kind words and zoom

sessions have helped me get through the last 4 years including this Covid-19 lockdown. To

Babybel my little companion, her constant company and cuddles got me through.

My grandparents, Pappou Nikitas and Yiayia Eleni who are no longer with us. Their foresight and

sacrifices in order to provide a better life for their family have enabled me to stand where I am

today. Pappou passed away during my candidature. He was president of my cheer-squad and

dreamed of attending my graduation with the ‘funny hat’ so that he could try it on too. He taught me

to have patience and to appreciate the educational opportunities I have been given. I hope that

wherever they are, Pappou and Yiayia are very proud that I have finished this PhD.

To my parents, Stavroula and Dusan, I dedicate this thesis to you. I cannot express how grateful I am

for your unwavering support throughout this journey. You are always there when I need you and you

knew when to step in even when I didn’t make it obvious that I needed support. Thank you for never

pressuring me to do anything, but always supporting me no matter which direction I chose to travel

– who knew we’d end up here! Thank you for your sacrifices which have given me the experiences

and opportunities I hold today. Mum, for all the times you came to stay with me, cook for me and

watch questionable reality TV with me. Dad, for always reminding me to have a work-life balance,

and a bag full of Aldi chocolates and food in the house. A massive thank you to both of you!

Lastly, I would like to thank the Australian Postgraduate Award and the Australian Government

Research Training Program Stipend Scholarship for their generous financial support with my research

scholarship.

VI

Contents Declaration ................................................................................................................... 2

Acknowledgements ..................................................................................................... III

List of Figures .............................................................................................................. X

List of Tables ............................................................................................................. XIV

Conferences & Awards ............................................................................................. XV

List of Abbreviations ................................................................................................ XVI

Summary ................................................................................................................... XIX

1 Introduction ........................................................................................................... 1

1.1 Intrauterine growth restriction (IUGR) ................................................................................... 1

1.2 Physiological impacts of IUGR ................................................................................................ 2

1.2.1 Impact of IUGR on growth and organ development ......................................................................... 2

1.2.2 Impact of IUGR on body composition .............................................................................................. 4

1.3 Impact of IUGR on the brain ................................................................................................... 5

1.4 Structure of the brain .............................................................................................................. 6

1.4.1 Structure of the cerebral hemispheres ............................................................................................... 6

1.4.2 Structure of the cerebellum ............................................................................................................. 10

1.5 Impact of IUGR on the brain .................................................................................................. 12

1.5.1 Impact of IUGR on the cerebrum .................................................................................................... 12

1.5.2 Impact of IUGR on the cerebellum ................................................................................................. 13

1.6 Animal models of IUGR used to assess brain injury............................................................... 14

1.7 Mechanisms of reduced and delayed myelination in IUGR .................................................... 15

1.8 Impact of IUGR on thyroid gland function ............................................................................ 18

1.8.1 Overview of TH signaling ............................................................................................................... 18

1.9 Role of TH in brain development ............................................................................................ 20

1.10 Use of thyroid hormone in neonatal brain injury ................................................................... 24

1.11 DITPA as a therapy in adults and children ............................................................................ 26

1.11.1 Physiological impacts of DITPA treatment. .................................................................................. 27

1.11.2 Impact of DITPA on the CNS and developing brain ..................................................................... 31

1.12 Scope of this thesis .................................................................................................................. 31

2 General Methodology ......................................................................................... 34

2.1 Introduction ............................................................................................................................ 34

2.2 Ethics clearance and animal welfare ....................................................................................... 35

2.3 Animals ................................................................................................................................... 35

2.4 Surgical procedure .................................................................................................................. 35

VII

2.4.1 Pre-operative preparation ................................................................................................................ 35

2.4.2 Surgery ............................................................................................................................................ 35

2.4.3 End of surgery and post-surgical care ............................................................................................. 37

2.5 Classification of IUGR and control, and size-matching of litters. ........................................... 37

2.6 Drug treatment ....................................................................................................................... 37

2.6.1 Handling of pups and injection........................................................................................................ 38

2.6.2 Monitoring after drug treatment ...................................................................................................... 38

2.7 Post-mortem blood and tissue collection ................................................................................. 38

2.7.1 Blood collection .............................................................................................................................. 39

2.7.2 Perfusion fixed brain tissue collection ............................................................................................ 39

2.7.3 Fresh snap frozen brain tissue collection ......................................................................................... 40

2.7.4 Processing of optic nerves ............................................................................................................... 40

2.8 Tissue histology ....................................................................................................................... 43

2.8.1 Paraffin sectioning of cerebral hemispheres and cerebellum .......................................................... 43

2.9 Histological staining & analysis .............................................................................................. 43

2.9.1 Haemotoxylin and Eosin (H&E) ..................................................................................................... 43

2.9.2 Analysis of H&E staining ................................................................................................................ 43

2.10 Immunohistochemical staining ............................................................................................... 43

3 Impact of DITPA treatment on myelination and inflammation in the

neonatal IUGR rat cerebrum ................................................................................... 46

3.1 Introduction ............................................................................................................................ 46

3.2 Methodology ........................................................................................................................... 49

3.2.1 Overview of animal work ................................................................................................................ 49

3.3 Paraffin sectioning of the cerebral hemispheres ..................................................................... 49

3.4 Immunohistochemical staining of the cerebral hemispheres .................................................. 50

3.5 Immunohistochemical analysis of the cerebral hemispheres .................................................. 50

3.5.1 Areal coverage (% AC) of MBP-, PLP- and GFAP-immunoreactivity (IR) ................................... 52

3.5.2 Projection of MBP- and PLP-IR fibres into the cerebral cortex ...................................................... 53

3.5.3 Areal density of Olig2-, APC- and Iba1-IR cells ............................................................................ 54

3.6 Statistical analysis ................................................................................................................... 55

3.7 Results ..................................................................................................................................... 57

3.7.1 Myelination and oligodendrocytes .................................................................................................. 57

3.7.2 Inflammation ................................................................................................................................... 69

3.8 Discussion ................................................................................................................................ 72

3.8.1 Overview ......................................................................................................................................... 72

3.8.2 Effects of daily DITPA administration on white matter development ............................................ 72

3.8.3 Effects of daily DITPA administration on inflammation in the cerebrum ...................................... 77

VIII

3.8.4 Limitations of the study ................................................................................................................... 77

3.8.5 Conclusion ....................................................................................................................................... 79

4 Impact of DITPA treatment on myelination and inflammation in the

neonatal IUGR rat cerebellum ................................................................................. 80

4.1 Preamble ................................................................................................................................. 80

4.2 Introduction ............................................................................................................................ 80

4.3 Methodology ........................................................................................................................... 83

4.3.1 Animals and tissue ........................................................................................................................... 83

4.3.2 Paraffin sectioning of the cerebellum .............................................................................................. 83

4.3.3 Assessment of the cerebellar structure ............................................................................................ 83

4.3.4 Immunohistochemical staining of the cerebellum ........................................................................... 84

4.3.5 Immunohistochemical analysis of the cerebellum .......................................................................... 85

4.3.6 Statistical analysis ........................................................................................................................... 88

4.4 Results ..................................................................................................................................... 91

4.4.1 Morphology of the cerebellum ........................................................................................................ 91

4.4.2 Immunohistochemical assessment of the cerebellum ...................................................................... 93

4.5 Discussion .............................................................................................................................. 101

4.5.1 Overview ....................................................................................................................................... 101

4.5.2 Effect of daily DITPA administration on cerebellar structure ...................................................... 101

4.5.3 Effect of daily DITPA administration on white matter development............................................ 102

4.5.4 Effect of DITPA administration on inflammation in the cerebellum. ........................................... 103

4.5.5 DITPA increased Purkinje cell linear density in early developing cerebellar lobules. ................. 105

4.5.6 Limitations of the study ................................................................................................................. 105

4.5.7 Conclusion ..................................................................................................................................... 106

5 Assessment of neonatal growth and wellbeing measures following DITPA

therapy in the IUGR rat. ........................................................................................ 107

5.1 Preamble ............................................................................................................................... 107

5.2 Introduction .......................................................................................................................... 107

5.3 Methodology ......................................................................................................................... 110

5.3.1 Overview of animal work .............................................................................................................. 110

5.3.2 Body and organ weights ................................................................................................................ 110

5.3.3 Analysis of body composition using dual-energy x-ray absorptiometry (DEXA) ........................ 111

5.3.4 Analysis of blood plasma .............................................................................................................. 112

5.3.5 Analysis of Dio1 in the Liver ........................................................................................................ 112

5.3.6 Statistical analysis ......................................................................................................................... 115

5.4 Results ................................................................................................................................... 116


IX

5.4.2 Brain weights ................................................................................................................................. 119

5.4.3 Body morphometry ........................................................................................................................ 123

5.4.4 Body composition (DEXA) ........................................................................................................... 125

5.4.5 Thyroid and liver function ............................................................................................................. 129

5.5 Discussion .............................................................................................................................. 134

5.5.1 Overview ....................................................................................................................................... 134


5.5.3 Brain weights ................................................................................................................................. 136

5.5.4 Body morphometry ........................................................................................................................ 136

5.5.5 Body composition (DEXA) ........................................................................................................... 137

5.5.6 Blood plasma analysis ................................................................................................................... 138

5.5.7 Limitations of the study ................................................................................................................. 140

5.5.8 Conclusion ..................................................................................................................................... 140

6 General Discussion ............................................................................................ 142

6.1 Overview ............................................................................................................................... 142

6.2 Does DITPA promote myelination in the cerebrum and therefore benefit the IUGR brain?

144

6.3 Is DITPA only beneficial when cerebral MCT8 is reduced? ................................................ 146

6.4 Should DITPA only be used in cases of confirmed IUGR? ................................................... 147

6.5 Future directions – clinical administration of DITPA .......................................................... 148

6.6 Conclusion ............................................................................................................................. 149

7 Reference List .................................................................................................... 150

Appendix 1 ............................................................................................................... 171

Appendix 2 ............................................................................................................... 172

Appendix 3 ............................................................................................................... 173

Appendix 4 ............................................................................................................... 175

Appendix 5 ............................................................................................................... 177

Appendix 6 ............................................................................................................... 180

Appendix 7 ............................................................................................................... 185

X

List of Figures

Figure 1.1 Diagrams showing the position of the hippocampus under the cerebral cortex, deep

within the medial temporal lobe ................................................................................................... 8

Figure 1.2 Diagram of a mid-sagittal cross-section through the human cerebellum, with the ten

lobules indicated by roman numerals (I-X). ............................................................................... 12

Figure 1.3 Timeline of oligodendrocyte development from pre-oligodendrocytes to the mature

myelinating phenotype in humans and rats. …………………………………………………...17

Figure 1.4 Thyroid hormone cellular signalling pathway. triiodothyronine, T4 = thyroxine, TR =

thyroid hormone receptor ............................................................................................................ 20

Figure 1.5 Timing of human and rat brain development in relation to thyroid hormone signalling..

..................................................................................................................................................... 22

Figure 1.6 Chemical structure of thyroid hormones and analogues .................................................. 27

Figure 2.1 Exposed rat uterus and uterine vessels during bilateral uterine artery ligation (BUVL)

surgery ......................................................................................................................................... 36

Figure 2.2 Experimental protocol timeline (A) and schematic diagram of tissue collection protocols

..................................................................................................................................................... 42

Figure 3. 1 Sequence of tissue sectioning and staining. .................................................................... 51

Figure 3.2 Coronal section of P14 rat cerebrum stained with MBP .................................................. 54

Figure 3.3 Areal coverage (% AC) of MBP-IR (A) in cortical layer VI, and proportion (%) of

cerebral cortex depth containing MBP-IR fibre projections (B) at P14 in control and IUGR

pups treated with DITPA or saline.. ............................................................................................ 58

Figure 3.4 Areal coverage (% AC) of MBP-IR in the corpus callosum (A), external capsule (B),

hippocampal CA1 region(C), hippocampal CA3 regions (D), and fimbria (E) at P14 in control

and IUGR pups treated with DITPA or saline. ........................................................................... 60

Figure 3.5 Percentage of area covered by PLP-IR (A) in the cortex (layer VI), and cortical

projection length (B) at P14 in control and IUGR pups treated with DITPA or saline. ............. 61

XI

Figure 3.6 Areal coverage (% AC) of PLP-IR in the corpus callosum (A, B), external capsule (C,

D), hippocampal CA1 (E) and CA3 (F) regions and fimbria (G) at P14 in control and IUGR

pups treated with DITPA or saline. ............................................................................................. 63

Figure 3.7 Areal density of Olig2-IR oligodendrocytes in the cortical layer VI (A), corpus callosum

(B), hippocampal CA1 region (C), hippocampal CA3 region (D), and fimbria (E) at P14 in

control and IUGR pups treated with DITPA or saline. ............................................................... 65

Figure 3.8 Areal density of APC-IR oligodendrocytes in the cortical layer VI (A), corpus callosum

(B), hippocampal CA1 region (C), hippocampal CA3 region (D), and fimbria (E) at P14 in

control and IUGR pups treated with DITPA or saline.. .............................................................. 67

Figure 3.9 Proportion of mature APC-IR OLs to total OL lineage (APC:Olig2) in the cortical layer

VI (A), corpus callosum (B), hippocampal CA1 region (C), hippocampal CA3 region (D), and

fimbria (E) at P14 in control and IUGR pups treated with DITPA or saline. ............................. 69

Figure 3.10 Density of Iba1-IR microglia in the cortex (layer VI; A), corpus callosum (B), CA1 (C)

and CA3 (D) regions of the hippocampus and fimbria (E) at P14 in control and IUGR pups

treated with DITPA or saline. ..................................................................................................... 70

Figure 3.11 Percentage of area covered by GFAP-IR astrocytes in the cortex (layer VI; A), corpus

callosum (B), external capsule (C), CA1 (D) and CA3 (E) regions of the hippocampus and

fimbria (F) at P14 in control and IUGR pups treated with DITPA or saline .............................. 71

Figure 4.1 H&E stained sagittal section of P14 rat cerebellum at the level of the vermis ................ 84

Figure 4. 2 Sequence of tissue sectioning and staining (A) A total of 40 sections were cut from each

tissue block; 8µm apart.. ............................................................................................................. 85

Figure 4.3 Olig2-immunostained sagittal section of P14 rat cerebellum at the level of the vermis

(counterstained with Haematoxylin). .......................................................................................... 87

Figure 4.4 Total cerebellar cross-sectional-area (A), layer widths (B – E) and layer width-to- total

cross-sectional-area (TCA; F – I) in control and IUGR pups treated with DITPA or saline at

P14 .............................................................................................................................................. 92

Figure 4.5 Width of the molecular layer in control and IUGR pups treated with DITPA or saline at

P14. ............................................................................................................................................. 93

Figure 4.6 Area coverage of MBP-IR in the cerebellar deep white matter (A) and lobule white

matter (B), and density of Olig2-IR oligodendrocytes in the deep white matter (C) and lobule

white matter (D) in control and IUGR pups treated with DITPA or saline at P14. .................... 94

XII

Figure 4.7 Cell density of Iba1-IR microglia in the cerebellar deep white matter (A) and lobule

white matter (B, C) in control and IUGR pups treated with DITPA or saline at P14.. .............. 95

Figure 4.8 Linear density of GFAP-IR Bergmann glia (BG) in the early (A) and late (B) developing

cerebellar lobules in control and IUGR pups treated with DITPA or saline. ............................. 97

Figure 4.9 Area coverage of GFAP-IR astrocytes in the cerebellar deep white matter (A), early and

late lobule white matter combined (B), as well as early lobules (C) and late lobules (D)

separately, in control and IUGR pups treated with DITPA or saline at P14. ............................. 98

Figure 4.10 Somal area (A), areal density (B) and linear density of calbindin-IR Purkinje cells in

cerebellar lobules combined (C), and in early (D) and late lobules (E) separately, in control and

IUGR pups treated with DITPA or saline at P14 ...................................................................... 100

Figure 5. 1 Overview of animals used in Chapter 5. A total of 223 P14 rat pups were used in this

study. This included both male and female control and IUGR pups, treated with DITPA or

saline. ........................................................................................................................................ 110

Figure 5. 2 Image of P14 rat taken using dual-energy x-ray absorptiometry (DEXA).. ................. 111

Figure 5.3 Body weights (g) at postnatal day 1 (A-C), P7 (D-F), and P14 (G-I) in control and IUGR

pups treated with DITPA or saline. ........................................................................................... 117

Figure 5.4 Liver (A-C) and kidney (D-I) weights (g) at P14 in male and female control and IUGR

pups treated with DITPA or saline. ........................................................................................... 119

Figure 5.5 Total brain weight (g) (A-C) and brain-to-body weight ratio (D-F) at P14 in male and

female control and IUGR pups treated with DITPA or saline. ................................................. 121

Figure 5.6: Weight (g) of the cerebral hemispheres (A-C), cerebellum (D-F), pons (G-I) and

medulla (J-L) in male and female control and IUGR pups at P14 treated with DITPA or saline.

................................................................................................................................................... 123

Figure 5.7: Crown-to-rump length (A-C), head circumference (D-F), and hip circumference (G-I)

(mm) in male and female control and IUGR pups at P14 treated with DITPA or saline. ........ 125

Figure 5.8 Bone mineral density (A-C), bone mineral content (D-F), and total bone area (G-I) in

male and female control and IUGR pups at P14 treated with DITPA or saline. ...................... 127

Figure 5.9 Lean tissue mass (A-C), total fat mass (D-F), and percentage (%) body fat (G-I) in male

and female control and IUGR pups at P14 treated with DITPA or saline. ............................... 129

XIII

Figure 5.10 FT3 (A-C) and FT4 (D-F) plasma levels in male and female control and IUGR pups at

P14 treated with DITPA or saline. ............................................................................................ 131

Figure 5.11 Serum levels of ALT (A-C), ALP (D-F), and cholesterol (G-I), and relative levels of

Dio1 to B2M (J; male liver only) in male and female control and IUGR pups at P14 treated

with DITPA or saline. ............................................................................................................... 133

Figure 5.12 Thyroid hormone homeostatic feedback system. ......................................................... 139

XIV

List of Tables

Table 1.1 Important DITPA studies in humans and animals. ............................................................ 30

Table 3.1 Summary of the 3 cerebral hemisphere levels at which analysis was carried out ............. 52

Table 3. 2 Immunohistochemistry - optimised antigen retrieval, blocking protocol and antibody

concentrations in the cerebrum. .................................................................................................. 56

Table 4. 1 Immunohistochemistry - optimised antigen retrieval, blocking protocol and antibody

concentrations in the cerebellum. ............................................................................................... 90

Table 5.1 Dio1 gene assay ID’s……………………………………………………………………………..114

Table 5. 2 Clinical reference ranges for plasma levels of free triiodothyronine (FT3) and free thyroxine

(FT4) in infants, children and adults. Reference ranges from Monash Pathology, Clayton, Vic,

Australia. ................................................................................................................................................ 131

Table 5.3 Clinical reference ranges for plasma levels of Alanine transaminase (ALT), alkaline phosphatase

(ALP) and cholesterol in infants, children and adults. Reference ranges from Monash Pathology,

Clayton, Vic, Australia. .......................................................................................................................... 133

Appendix 3, Table 3. 1 Two-way ANOVA results of MBP-IR analysis, supplementary to Chapter

3, Section 3.7.1. ......................................................................................................................... 173

Appendix 3, Table 3. 2 Two-way ANOVA results of PLP-IR analysis, supplementary to Chapter 3,

Section 3.7.1. ............................................................................................................................. 173

Appendix 3, Table 3. 3 Two-way ANOVA results of Olig2–IR analysis, supplementary to Chapter

3, Section 3.7.1.. ........................................................................................................................ 174

Appendix 3, Table 3. 4 Two-way ANOVA results of APC–IR analysis, supplementary to Chapter

3, Section 3.7.1.. ........................................................................................................................ 174

Appendix 3, Table 3. 5 Two-way ANOVA results of APC-IR: Olig2-IR analysis, supplementary to

Chapter 3, Section 3.7.1. ........................................................................................................... 174

Appendix 4, Table 4. 1 Two-way ANOVA results of cerebellar morphology measurements using

H&E staining supplementary to Chapter 4 Section 4.4.1 ......................................................... 175

Appendix 4, Table 4. 2 Two-way ANOVA results of immunohistochemical analysis in the

cerebellum, supplementary to Chapter 4 Section 4.4.2. ........................................................... 175

XV

Conferences & Awards

International Conferences

• 3rd Congress of joint European Neonatal Societies (jENS) 2019, Maastricht, Netherlands,

Poster presentation: Kondos-Devcic, D., Wong, F., Cumberland, C., Khore, M., Walker, D.,

Tolcos, M. “Assessment of neonatal growth and wellbeing following thyroid hormone based

therapy in a rodent model of intrauterine growth restriction (IUGR)”.

• International Society for Developmental Origins of Health and Disease (DOHaD) World

Congress 2019, Melbourne, Australia, Poster presentation: Kondos-Devcic, D., Wong, F.,

Cumberland, C., Khore, M., Walker, D., Tolcos, M. “Assessment of neonatal growth and

wellbeing following thyroid hormone based therapy in a rodent model of intrauterine growth

restriction (IUGR)”.

National Conferences

• 33rd Annual Fetal and Neonatal Workshop of Australia and New Zealand 2019, as part of the

23rd Congress of the Perinatal Society of Australia and New Zealand (PSANZ) 2019, Surfers

Paradise, Queensland. Oral presentation: Kondos-Devcic, D., Wong, F., Cumberland, C.,

Khore, M., Walker, D., Tolcos, M. “Assessment of neonatal growth and wellbeing following

thyroid hormone-based therapy in a rodent model of intrauterine growth restriction (IUGR)”.

• 31st Annual Fetal and Neonatal Workshop of Australia and New Zealand 2019, Canberra,

Australian Capital Territory. As part of the 21st Congress of the Perinatal Society of Australia

and New Zealand (PSANZ) 2017, Canberra, Australian Capital Territory.

• HDR Student Symposium 2019, RMIT Bundoora Campus, Victoria. Oral presentation: “A

Thyroid hormone-based treatment to restore the brain maturation following foetal growth

restriction”.

• Students of Brain Research (SOBR) Conference 2018, St Vincent’s Hospital, Melbourne,

Victoria. Kondos-Devcic, D., Walker, D., Wong, F., Tolcos, M. Oral presentation: “A

Thyroid hormone-based treatment to restore the brain in growth restricted babies”.

Awards

1. RMIT University Bundoora Campus Joint 2nd Place, 3-Minute Thesis competition 2018.

2. RMIT HDR Beautiful Science Competition 2019 1st Prize for scientific image.

3. Australian Postgraduate Award, 2016.

XVI

List of Abbreviations

% AC

<

±

1°

2°

*

ABC

AHDS

ALP

ALT

ANOVA

APC

ATP

BG

BMC

BMD

BSA

BUVL

CA1

CA3

cm2

CNS

CRL

DAB

DEXA

dg

dH2O

DITPA

Dio1

DNA

DPX

DWM

E

EGL

F

FT3

FT4

fMRI

Percentage of areal coverage

Less than

Plus/minus

Primary

Secondary

Multiplication sign

Avidin/Biotinylated enzyme Complex

Allan-Hernon-Dudley syndrome

Alkaline phosphatase

Alanine transaminase

Analysis of variance

Adenomatous polyposis coli

Adenosine triphosphate

Bergmann glia

Bone mineral content

Bone mineral density

Bovine serum albumin

Bilateral uterine vessel ligation

Cornu ammonis region 1

Cornu ammonis region 3

Centimetres squared

Central nervous system

Crown-to-rump length

3,3’- Diaminobenzadine

Dual energy x-ray absorptiometry

Days of gestation

Distilled water

3,5-diiodothyropropionic acid

Deiodinase 1

Deoxyribonucleic acid

Dibutylphthalate polystyrene xylene

Deep white matter

Embryonic day

External granular layer

Ratio of residual variances

Free plasma 3,5,3’- triiodothyronine

Free plasma thyroxine

Functional magnetic resonance imaging

XVII

GA

GABA

GFAP

GM

H2O2

H&E

Iba1

IGL

i.p.

IR

IUGR

IU/L

IQ

KO

MBP

MCT8

ML

MRI

mRNA

NaCl

NHMRC

N

OL/s

Olig2

OPCs

P

p

PB

PBS

PCR

PFA

PLP

RNA

ROI

SD

SEM

SVZ

T3

T4

TCA

TH

TSH

Gestational age

Gamma-Aminobutyric acid

Glial fibrillary acidic protein

Grey matter

Hydrogen peroxide

Haemotoxylin and Eosin

Ionized calcium-binding adaptor molecule 1

Internal granular layer

Intraperitoneal

Immunoreactivity/ immunoreactive

Intrauterine growth restriction

International units per litre

Intelligence quotient

Knockout

Myelin basic protein

Monocarboxylate transporter-8

Molecular layer

Magnetic resonance imaging

Messenger ribonucleic acid

Sodium chloride

National Health and Medical Research Council

Sample size

Oligodendrocyte/s

Oligodendrocyte transcription factor 2

Oligodendrocyte progenitor cells

Postnatal day

p-value

Phosphate buffered

Phosphate buffered saline

Polymerase chain reaction

Paraformaldehyde

Myelin proteolipid protein

Ribonucleic acid

Region of interest

Standard deviation

Standard error of the mean

Subventricular zone

3,5,3’- triiodothyronine

3,5,3’,5’ – tetraiodothyronine or thyroxine

Total cross-sectional cerebellar area

Thyroid hormone

Thyroid-stimulating hormone

XVIII

TR

TRH

USA

WM

Thyroid hormone receptor

Thyrotropin-releasing hormone

United States of America

White matter

Summary

XIX

Summary

Intrauterine growth restriction (IUGR) is a condition in which a foetus does not reach its full genetic

growth potential, and is a leading cause of perinatal death and postnatal morbidity. IUGR often results

in permanent neurodevelopmental deficits, ranging from cognitive and behavioural impairments to

cerebral palsy. IUGR can occur due to a number of environmental or genetic factors, however the

dominant cause is a lack of oxygen and nutrients delivered the foetus as a result of placental

insufficiency. Deficits in grey matter (GM) and white matter (WM) in the foetal brain are thought to

underlie these neurodevelopmental sequelae. There is no current treatment to prevent or correct

placental insufficiency, and babies are often delivered preterm if IUGR is found to be severe or

worsening, further increasing the potential for adverse outcomes. With 27,000 IUGR babies born in

Australia in the past year alone, and 30 million born worldwide, new therapies which can be delivered

immediately after birth, are urgently required.

IUGR decreases the expression of monocarboxylate transporter-8 (MCT8) (Azhan, 2019, Chan et al.,

2014), a transporter protein necessary for the delivery of thyroid hormone (TH) into cells such as

oligodendrocytes (OLs) in the brain (Lee et al., 2017). TH is critical for the maturation of OLs during

foetal brain development, and reduced transport of TH into OLs impairs myelination (Lee et al.,

2017). In this thesis, it is proposed that the synthetic TH analogue 3,5-diiodothyropropionic acid

(DITPA) which does not require MCT8 to enter cells, can be used to overcome deficits caused by a

loss of MCT8 expression in the IUGR brain, thereby restoring myelination. Our group has previously

shown that MCT8 mRNA levels are reduced in the newborn IUGR rat at postnatal day (P) P7, and

these levels normalise by P14 (Azhan, A., PhD thesis, Monash University, 2019). Furthermore, we

have previously shown that short-term administration of DITPA (0.5mg/100g/day i.p.) to newborn

IUGR rats (from P1 to P6) corrected the myelination deficit in the external capsule by P7, and did

not affect neonatal growth parameters (Azhan, A., PhD thesis, Monash University, 2019). The present

project set out to investigate the benefits of a longer-term DITPA treatment from P1 to P13

(0.5mg/100g/day i.p.) a time in rat brain development equivalent to 23 to 40 weeks gestation in the

human (Semple et al., 2013), and reflective of what is likely to occur in the clinical setting with an

IUGR baby delivered preterm and given DITPA until term equivalent age. The same cohort of

animals contributed to all 3 experimental chapters in this thesis.

In Chapter 3, the impact of DITPA administration on the development of the cerebrum was

investigated, focusing on myelination, a neurodevelopmental process affected in IUGR, and regions

Summary

XX

with known vulnerability to the prenatal insults. Specifically, this study aimed to determine whether

DITPA administration in IUGR rat pups from P1 to P13 improved myelination, promoted OL

maturation, and did not cause injury or inflammation in the cerebrum compared to vehicle (saline)

treatment. The data presented in Chapter 3 shows that DITPA treatment in IUGR may promote

myelination in the cerebral cortex and fimbria, increase OL density in the corpus callosum, and does

not cause injury or inflammation in the cortex, corpus callosum, hippocampus or fimbria when

assessed at P14. This study indicates that an extended duration of DITPA treatment may be beneficial

to myelination in the IUGR cerebrum but is not favourable when given in controls, decreasing myelin

proteolipid protein (PLP) and OLs the cortex and fimbria. Next, it was essential to investigate

DITPA’s therapeutic potential in another brain region with known vulnerability to IUGR (De Bie et

al., 2011, Padilla et al., 2011), the cerebellum.

The study presented in Chapter 4 aimed to determine the impact of DITPA treatment (P1 to P13) on

myelination, OL development, morphology, and neuroinflammation in the cerebellum in IUGR rat

pups. DITPA did not improve myelination or promote OL maturation in the cerebellum of IUGR

pups when assessed at P14, had no negative impact on cerebellar morphology (layer widths/areas and

Bergmann glial fibre density), however increased the density of Purkinje cells, and microglia in late

developing cerebellar lobules. Overall, these data support longer-term DITPA administration in

IUGR as exhibiting more benefit in the cerebrum than the cerebellum; as in the cerebrum DITPA

may be unfavourable when given to controls. To further investigate the potential use of DITPA in

treating IUGR, potential off-target effects on neonatal growth and wellbeing were determined next.

Chapter 5 examined potential off-target effects of DITPA administration in control and IUGR

neonates, focussing on neonatal growth, and assessment of wellbeing. Body weight of all pups was

assessed at P1, P7 and P14. Liver and kidney weights, body morphometry (head and hip

circumference and crown-to-rump length), body composition (bone density and mineral content, lean

tissue mass, fat mass), thyroid and liver function, as well as cholesterol levels were measure in pups

at post-mortem (P14). DITPA administration in IUGR pups did not adversely impact neonatal growth,

brain or organ weights or body composition, despite altering FT4 levels and showing hepatic

thyromimetic activity, but alters growth, liver weight and bone mineral content when given to control

pups.

In conclusion, this thesis has, for the first time, demonstrated that longer-term administration of the

TH analogue DITPA, which enters cells independently of MCT8, to newborn IUGR rats promotes

myelination in the cerebrum, increases OL density in the corpus callosum, and does not cause injury

Summary

XXI

or inflammation to the brain, albeit for a possible inflammatory response in the late developing

cerebellar lobules. In IUGR pups there were no negative off-target effects on neonatal growth or

wellbeing, although DITPA caused unfavourable outcomes when administered to controls. The work

presented in this thesis collectively highlights the potential for DITPA to improve myelination

outcomes in the IUGR brain, without causing injury or adverse off-target physiological effects,

however further studies are required before DITPA can be considered as a therapy in IUGR.

Introduction

1

1 Introduction

1.1 Intrauterine growth restriction (IUGR)

Intrauterine growth restriction, or IUGR is characterised by foetal growth that is small for gestational

age (GA) due to environmental or genetic factors, and is a major clinical challenge (Murki, 2014). In

developed countries, up to 9% of all pregnancies are affected by IUGR; in Australia this equates to

27,000 born in the past year, and 30 million infants born worldwide (de Onis et al., 1998, Wardlaw,

2004). IUGR is second to prematurity as the leading cause of perinatal morbidity and mortality (Abu-

Saad and Fraser, 2010, Bhutta et al., 2005), with IUGR babies at an increased risk of adverse

neurodevelopmental sequelae (Geva et al., 2006b), and 10 – 30 fold increased risk of developing

cerebral palsy (McIntyre et al., 2013, MacLennan et al., 2015). Nearly a third of IUGR births are due

to genetic causes, with the rest related to the foetal environment (Murki, 2014), and largely due to

deficient oxygen and nutrient supply to the foetus caused by maternal, placental or foetal factors.

IUGR can be a consequence of damage to, or partial occlusion of the umbilical cord (Murki, 2014,

Nash and Persaud, 1988), as well as placental malfunction (Krishna and Bhalerao, 2011). The most

common cause of IUGR is placental insufficiency, characterised by compromised utero-placental

blood flow, attributed to developmental faults in the placenta or placental blood vessels, thought to

be due to altered expression of growth factors (Regnault et al., 2002). This results in the foetus

receiving less blood, and therefore less oxygen and nutrients (Gaccioli and Lager, 2016). If serious,

IUGR is detected during late pregnancy, and the baby is often delivered preterm in order to remove

it from the insufficient intrauterine environment.

IUGR foetuses and neonates are classified as having symmetric, asymmetric or mixed growth

restriction, dependent on the ratio of head to abdominal circumference (foetus) or growth percentiles

of head circumference versus weight and length (neonate). IUGR commencing early in gestation

(prior to 32 weeks of gestation) and associated with symmetrical growth restriction, is more

commonly associated with chromosomal abnormalities or congenital infections (Sharma et al.,

2016a). Later-onset IUGR on the other hand, is usually associated with asymmetrical IUGR and is

often a result of placental disorders (Rosenberg, 2008). Asymmetrical IUGR infants are characterised

by a normal head circumference, with a proportionally smaller body weight, commonly due to “brain

sparing” mechanisms that occur during IUGR (Flood et al., 2014). Lastly, mixed IUGR occurs when

pre-existing growth restriction is affected further by placental disorders (Sharma et al., 2016b). The

Introduction

2

IUGR foetus often displays an adaptive mechanism, known as the ‘foetal brain-sparing effect’,

whereby in response to placental insufficiency, there is vasodilatation of the foetal cerebral circulation

to protect the brain. However this typically occurs at the expense of body weight and of other organs

like the kidneys and liver, which remain growth restricted (Miller et al., 2016). This thesis will focus

on IUGR caused by placental insufficiency, as it is the most common clinical presentation.

1.2 Physiological impacts of IUGR

Reduced oxygen and nutrient availability to IUGR foetuses as a result of placental insufficiency result

in a number of unfavourable physiological outcomes. While the defining characteristic of IUGR

neonates is their reduced body weight, there are also many associated physiological effects including

hypoglycaemia (Cowett et al., 1984, Haymond et al., 1974), altered blood cell counts (Castle et al.,

1986, Van den Hof and Nicolaides, 1990, Snijders et al., 1993), impaired organ growth (Platz and

Newman, 2008, Man et al., 2016) and body composition measures (Namgung and Tsang, 2000,

Namgung and Tsang, 2003, Chunga Vega et al., 1996) and altered thyroid gland activity (Kilby et al.,

2000, Soothill et al., 1992, Thorpe-Beeston et al., 1991). Reduced lipid and glycogen storage is seen

in IUGR foetuses as a result of placental insufficiency, and this contributes to neonatal

hypoglycaemia, where blood sugar levels are reduced (Cowett et al., 1984, Haymond et al., 1974).

IUGR neonates also display reduced white blood cell counts (leucopenia), as well as reduced platelet

numbers (thrombocytopenia) (Castle et al., 1986), both essential for immune defence. For the

purposes of this thesis, a focus will be placed on the impact of IUGR on growth of the body and

organs such as the kidneys and liver, and body composition measures including bone mineral density

and content, lean tissue mass and fat mass, as well as thyroid function and liver activation. These

parameters are all known to be impacted by IUGR (Bernstein et al., 2000, Martorell et al., 1998, Pena

et al., 1988, Strauss and Dietz, 1998, Kilby et al., 1998, Soothill et al., 1992, Thorpe-Beeston et al.,

1991).

1.2.1 Impact of IUGR on growth and organ development

While approximately 70 to 90% of IUGR infants display some form of catch-up growth from birth to

two years of age, they often fail to catch up completely (Monset-Couchard and de Bethmann, 2000).

In IUGR-born children, reduced growth and development is seen at 12 months of age (Low et al.,

1978), and reduced height is seen at school age, with a reduction in head circumference compared to

children who were born with appropriate weight for GA (Robertson et al., 1990, Monset-Couchard

et al., 2004). Overall, IUGR neonates maintain a smaller stature in adulthood compared to adults who

were born appropriately sized (Karlberg and Albertsson-Wikland, 1995, Westwood et al., 1983). This

is also seen in animal models of IUGR, where reduced body weight and crown-to-rump length are

Introduction

3

observed in IUGR rats at P7 and P14 compared to non-IUGR born rats (McDougall et al., 2017b,

1984, Olivier et al., 2005, Wlodek et al., 2005), and in IUGR foetal guinea pigs (Herrera et al., 2016)

at 52 days of gestation (dg), 60 dg, 1 week, and 8 weeks postnatal age (Tolcos et al., 2018, Tolcos et

al., 2011). It is important to note, that experimental models of IUGR such as bilateral uterine vessel

ligation, do not produce offspring that are equally IUGR. Instead there is a spectrum of growth

restriction that is dependant on the positioning of the foetus relative to the ligation site. Of relevance

to this thesis, it has been well documented in rats that foetuses in the uterine horn which are positioned

closest to the site of ligation generally experience the highest level of growth restriction compared to

their littermates and are more likely to die in utero resulting in a smaller litter size. Foetuses further

away from the ligation site may benefit more from supplementary blood supply(Hayashi and Dorko,

1988, Wigglesworth, 1964, Wigglesworth, 1974, Catteau et al., 2011, Gallo et al., 2012) This

experimental ‘range’ of IUGR in the rat should be taken into account when discussing outcomes.

IUGR also affects organ growth, where compared to non-IUGR counterparts, IUGR infants have

significantly smaller brains, hearts, lungs, thyroid glands, livers and kidneys (Platz and Newman,

2008, Man et al., 2016). This thesis will assess the impact of IUGR on liver and kidney growth, two

organs known to be vulnerable to the effects of IUGR (Schreuder et al., 2006). The liver is essential

for detoxification, metabolism and immune function, as well as synthesising proteins, metabolites

and biochemical enzymes that are essential for digestion and metabolism in the body. Relevant to this

thesis, the liver metabolises drugs (Remmer, 1970), as well as regulating energy by storing excess

blood glucose as glycogen upon being stimulated with insulin, which is released by the pancreas, and

then converting it back to glucose when the body requires energy. In humans, IUGR is associated

with insulin resistance in childhood (Veening et al., 2002, Hofman et al., 1997, Fraser et al., 2007)

and adulthood (Flanagan et al., 2000, Jaquet et al., 2000), as well as the prevalence of metabolic

disorders, the presentation of ‘fatty liver disease’ (Alisi et al., 2011), and abnormal glucose tolerance

in adulthood (Hales et al., 1991, Lithell et al., 1996). These studies highlight that impairments in liver

function as a result of IUGR are long lasting. Animal models provide further insight, with impaired

insulin signalling, as well as reduced amino acid uptake seen in the foetal IUGR sheep liver (Thorn

et al., 2009), while reduced glycogen storage (Oh et al., 1970), and decreased liver glucose

transporters are seen in IUGR rats compared to controls (GLUT-1 & GLUT-2) (Lane et al., 2001a).

In IUGR foetal pigs, lipoprotein lipase an important protein for the metabolism of vitamins, minerals,

protein and importantly glucose is reduced (Liu et al., 2013), suggesting that IUGR hinders the liver’s

ability to extract nutrients. IUGR also markedly reduces liver weight, with reduced liver weight in

human IUGR foetuses, being indicative of functional disturbances (Latini et al., 2004, Ladinig et al.,

2014, Molina Giraldo et al., 2019). Reduced liver weight is also seen in IUGR rats (Azhan, A., PhD

Introduction

4

thesis, Monash University, 2019) and sheep (Limesand et al., 2006). It is believed that the level of

organ ‘catch-up growth’ following IUGR underpins the impaired metabolic outcomes seen (Singhal

et al., 2003). Together, these studies suggest that impaired liver function following IUGR is likely to

be correlated with decreased liver weight and level of catch-up growth. IUGR also reduces foetal

circulating cholesterol levels (Pecks et al., 2012, Pecks et al., 2016, Bon et al., 2007, Merzouk et al.,

1998, Roberts et al., 1999). Cholesterol is and is a lipid-rich substance that is essential for cellular

structure, and is synthesised by the liver, or ingested through diet, however when present in high

amounts can increase the risk of cardiac disease. IUGR is shown clinically to predispose adults to

elevated circulating cholesterol levels (Barker et al., 1993).

The kidneys (renal system) are the body’s filtration system containing nephrons which act as filtration

units to remove waste and extra fluid in urine. The kidneys are responsible for maintaining a

homeostatic balance of electrolytes, minerals and water within the body, and in IUGR foetuses as

well as during the first year of life and into childhood (Naeye, 1965, Hotoura et al., 2005, Schmidt et

al., 2005, Latini et al., 2004), kidney volume is reduced compared to controls, indicative of decreased

nephron density (Silver et al., 2003). Indeed, decreased nephron density has long been correlated with

IUGR (Brenner and Chertow, 1993), and this pathology is also associated with outcomes of renal

dysfunction and hypertension later in life and in humans (Chan et al., 2010) and rats (Battista et al.,

2002) (R. Valdez, 1994, M. do Carmo Pinho Franco, 2003, Mackenzie and Brenner, 1995). These

studies collectively highlight the negative implications of IUGR on the liver, cholesterol levels and

renal system, and suggest that IUGR predisposes individuals to negative health outcomes later in life.

1.2.2 Impact of IUGR on body composition

IUGR impacts body composition measures such as bone mineral density and content, lean tissue mass

such as muscle, as well as body fat. Clinically, IUGR foetuses have a 25 to 40 % decrease in skeletal

muscle mass compared to their non-IUGR foetuses, when assessed using ultrasound in late gestation

(Larciprete et al., 2005). Bone mineral density and content is also reduced in the IUGR foetus

(Verkauskiene et al., 2007), and at birth (Namgung and Tsang, 2000, Namgung and Tsang, 2003,

Chunga Vega et al., 1996). Lean tissue mass and muscle mass is reduced in IUGR compared to non-

IUGR counterparts (Srikanthan and Karlamangla, 2011), and this persists postnatally (Beltrand et al.,

2008, Larciprete et al., 2005), with reduced muscle mass and impaired physical strength extending

into adulthood (Brown and Hay, 2016). A lower percentage of body fat is observed in IUGR foetuses

(Larciprete et al., 2005), and infants (Verkauskiene et al., 2007) compared to those who are not growth

restricted. Animal models of IUGR (induced via placental insufficiency) display similar findings,

with reduced skeletal mass reported in foetal sheep compared to controls (Costello et al., 2008), and

Introduction

5

reduced bone mineral concentration found in IUGR rats at P7 (Azhan, A., PhD thesis, Monash

University, 2019). Muscle mass is significantly reduced in IUGR lambs compared to controls (Fahey

et al., 2005), and reduced lean tissue mass and body fat in IUGR rats at P7 (Azhan, A., PhD thesis,

Monash University, 2019) and at 5 months of age (Coupe et al., 2012). It is clear from the numerous

studies across species that IUGR negatively impacts body composition measures, however IUGR is

also known to disrupt brain development (Batalle et al., 2012, Lodygensky et al., 2008, Tolsa et al.,

2004), as discussed below.

1.3 Impact of IUGR on the brain

The brain is the most complex organ in the human body, containing approximately 15 to 33 billion

neurons, which transmit afferent and efferent information (Herculano-Houzel, 2009). Apart from

being essential for cognition, the brain orchestrates the correct functioning of organs, muscles and

endocrine activity, allowing rapid and coordinated responses to changes in environment. The brain is

comprised of three main compartments, the forebrain (cerebrum) which contains the cerebral cortex,

the midbrain and the hindbrain, which contains the cerebellum. These regions are comprised of GM,

regions that are dense in neurons, axons and neuroglia, and WM, which contain myelinated axons

and neuroglia. The focus of this thesis will be on the cerebrum, including the cerebral cortex and

subcortical structures like the corpus callosum, external capsule and hippocampus, as well as on the

cerebellum, as these regions of the brain are known to be highly vulnerable to prenatal insults like

IUGR, often resulting in long-term neurological impairment (Padilla et al., 2011, Padilla et al., 2014b,

Egana-Ugrinovic et al., 2014, Lodygensky et al., 2008). Surviving IUGR infants have a greatly

increased risk of neurodevelopmental impairment (Geva et al., 2006b), which range from learning

difficulties (Geva et al., 2006a), decreased intellectual and cognitive function (de Bie et al., 2010,

Geva et al., 2006a) to a 10 to 30 fold increased risk of developing cerebral palsy (McIntyre et al.,

2013, MacLennan et al., 2015). Reduced regional and total brain volumes have been found in

foetuses, neonates, children and adolescents who were born IUGR (Padilla et al., 2014b, Tolsa et al.,

2004, Dubois et al., 2008). This is thought to be largely due to loss of GM and microstructural changes

in the WM, suggesting reduced myelination and axonal injury (Samuelsen et al., 2007, Padilla et al.,

2011, Padilla et al., 2014b, Businelli et al., 2014, Tolsa et al., 2004, Dubois et al., 2008, Batalle et al.,

2012, Eikenes et al., 2012a, Egana-Ugrinovic et al., 2014). Post-mortem studies show disrupted

myelination in the brain of preterm IUGR infants (Chase et al., 1972), while in vivo neuroimaging

studies show that disruption to myelin integrity can persist to adulthood (Padilla et al., 2014a, Esteban

et al., 2010, Eikenes et al., 2012b). Before the impact of IUGR on the brain is discussed in further

detail, a brief summary of brain development and structure of the mature brain will be provided to

enable greater understanding of the studies within this thesis.

Introduction

6

1.4 Structure of the brain

Development of the central nervous system (CNS) and brain begins as early as 4 weeks GA, where

following gastrulation, ectodermal specification and closure of the neural tube, neuronal progenitors

begin differentiating to form three vesicles; the prosencephalon, mesencephalon and

rhombencephalon (the developing forebrain, midbrain and hindbrain brain respectively). The

prosencephalon, or forebrain subsequently differentiates into 2 separate vesicles known as the

telencephalon, which later forms the cerebral hemispheres, and the diencephalon. The

rhombencephalon or hindbrain also further divides into 2 vesicles known as the metencephalon,

which later forms the cerebellum and pons, and the myelencephalon which gives rise to the medulla.

This thesis will focus on the cerebral hemispheres (including the cerebral cortex, corpus callosum,

external capsule and hippocampus) as well as the cerebellum, with emphasis placed on cerebellar

layer morphology and myelination. Where known, the developmental time-points discussed below

(Sections 1.4.1 & 1.4.2) will refer to the rat/rodents as embryonic age (E) or postnatal day (P).

1.4.1 Structure of the cerebral hemispheres

During the 10th week of gestation, growth of the brain hemispheres gives rise to the anterior frontal

lobes, and the posterior parietal, occipital and temporal lobes of the cerebrum. Cerebral development

can be divided into the early embryonic and late foetal phases. During the early embryonic phase

which occurs during the 5th to 8th week GA (Marin-Padilla, 1983), the cerebral vesicles begin to form,

and GM that will later become the mature six-layered cerebral cortex begins to stratify and

differentiate (Marin-Padilla, 1983). The mature cerebrum consists of two hemispheres, which contain

four lobes - the frontal, parietal, occipital and temporal lobes - which are responsible for motor,

sensory, visual and auditory functioning (Shi et al., 2012). Development of the six-layered cerebral

cortex, as well as the corpus callosum, external capsule and hippocampus will be outlined in greater

detail below.

1.4.1.1 Structure of the cerebral cortex

Development of the cerebral cortex begins with the differentiation of neurons from cortical stem cells

and progenitor cells within a region known as the ventricular zone. This is followed by an extended

period of neurogenesis resulting in mature functional neurons with electrophysiological properties

and the ability to form functional excitatory synaptic networks (Shi et al., 2012). Neuronal

proliferation consists of ventricular stem cells and progenitor cells undergoing a number of highly

regulated steps as they migrate dorsally to take their position in the six-layered morphology of the

mature cerebral cortex (Noctor et al., 2004, Wonders and Anderson, 2006). The cerebral cortex is

Introduction

7

generated in an ‘inside-out’ manner, with deep layer neurons being produced first, and more

superficial layer neurons arising last. In order of formation from deepest to most superficial the layers

are: layer VI (multiform layer), V (internal pyramidal layer), IV (internal granular layer), III (external

pyramidal layer), II (external granular layer) and I (molecular layer). Layers V and VI combined are

known as the infragranular layer and connects the cortex with subcortical regions. Layer V contains

stellate cells and gives rise to efferent projections to the basal ganglia, brainstem and spinal cord,

while layer VI contains fusiform cells, primarily projecting to the thalamus. Once fully developed,

the cerebral cortex is comprised of tightly folded GM, and contains the six-layered neocortex

(Rhoton, 2002), which contains excitatory and inhibitory neurons, glial cells such as astrocytes

(Luskin et al., 1993), microglia and OLs (Harris et al., 2015), and endothelial cells which constitute

blood vessels (Garcia-Cabezas et al., 2016). Between 16 to 18 weeks GA in humans, a wave of

oligodendrocyte progenitor cells (OPCs) migrate dorsally from a region known as the medial

ganglionic eminence, and populate the developing cortex (Kessaris et al., 2006, Rakic S, 2002). This

occurs from E15.5 in the mouse brain (Miller, 2002). A final wave of OPCs arise postnatally in

humans, and at birth in mice; the timing of this is mediated by the gene EMX1 being expressed by

cells in the ventricular zone of the developing cerebral cerebrum (Gorski et al., 2002, Winkler et al.,

2018). Development of OLs continue throughout childhood, adolescence and into adulthood

(McKenzie et al., 2014, Miller, 2002, Richardson et al., 2006). Beneath the cerebral cortex lies the

subcortical WM, and structures including the corpus callosum, external capsule and hippocampus

which will be a focus of this thesis and discussed below.

1.4.1.2 Structure of the hippocampus

The hippocampus develops in the medial temporal lobule, beneath the cerebral cortex, and is

responsible for navigation and spatial memory. Neuronal development and myelination in the human

and rodent hippocampus proceeds throughout childhood and into adulthood (Arnold and

Trojanowski, 1996, Gould et al., 1999, Eriksson et al., 1998, Stanfield and Trice, 1988, Cameron et

al., 1993). By 15 to 19 weeks GA the regions of the hippocampus are distinguishable, and from 20 to

25 weeks GA it expands substantially in volume (Arnold and Trojanowski, 1996). The region of the

hippocampus which later develops into the dentate gyrus, wraps around what will later become the

cornu ammonis region 3 (CA3), consisting of densely packed immature granule neurons. From 39 to

40 weeks (term) the hippocampus is mature in structure in humans, and at ~ P22 to 24 is mature in

rodents (Charvet and Finlay, 2018). Myelination in the hippocampus first appears at ~ 39 weeks GA

in humans (Arnold and Trojanowski, 1996), and at ~ P22 to 23 in rodents (Soriano et al., 1994). OLs

are first seen in the fimbria (fimbriae plural), a bundle of afferent and efferent nerve fibres adjacent

to the hippocampus proper at 20 weeks GA in humans (Abraham et al., 2010). The structure of the

Introduction

8

hippocampus is mature at birth, consisting of an adjacent cortical structure known as the hippocampal

gyrus, and a strip of densely packed neurons positioned between the hippocampus and the

hippocampal gyrus, called the dentate gyrus. The hippocampal gyrus consists of the entorhinal cortex

and subiculum, both involved in propagating the flow of information through the hippocampus

(Duvernoy, 2005) (Figure 1.1 C). Information is received from the overlying cerebral cortex via the

entorhinal cortex, projecting to the dentate gyrus (Duvernoy, 2005). Axon fibres then leave the

dentate gyrus and project to the cornu ammonis region (CA3), a region characterised by neuronal

morphology which is not uniform compared to the other hippocampal regions, and which plays an

essential role in encoding short-term memory and spatial information (Cherubini and Miles, 2015).

The CA3 propagates information to the cornu ammonis region 1 (CA1), which contains smaller

neurons than the CA3, and is integral for encoding long-term memory and spatial recognition. The

CA1 then relays impulses to neurons in a region called the subiculum which projects back to the

enterohinal cortex, enabling the signals to extend to the rest of the brain. Impulses can also leave the

hippocampus through the fimbria, subsequently entering the fornix, a WM fibre bundle (Figure 1.1

B) which connects the hippocampus to a variety of subcortical structures like the thalamus and

hypothalamus (Duvernoy, 2005).

Figure 1.1 Diagrams showing (A) the position of the hippocampus under the cerebral cortex, deep within the

medial temporal lobe, (B) the structure of the fornix and fimbriae which connects the hippocampus to

subcortical structures and (C) the structure of mature hippocampal formation including the dentate gyrus

(pink), cornu ammonis (yellow), entorhinal cortex (green) and parahippocampal gyrus (blue).

Introduction

9

1.4.1.3 Structure of the corpus callosum

The corpus callosum is a large bundle of myelinated neuronal fibres which transverses the midline of

the brain, providing connectivity between the left and right cerebral hemispheres through intra, as

well as inter-hemispheric axonal projections (Schulte and Muller-Oehring, 2010), that connect the

cortex in the frontal, parietal, occipital and temporal lobes of the brain (Aralasmak et al., 2006). The

corpus callosum is the largest WM tract in the human brain (Fitsiori et al., 2011), and rat brain

(Olivares et al., 2001), and contains sub-regions including the genu, midbody, truncus and splenium

which connect the corpus callosum to sensory, auditory, motor and somatosensory regions within the

brain (Goldstein et al., 2020). The structure of the corpus callosum in humans and rodents is similar,

and development, including myelination continues into adulthood in both humans and rats (Nunez et

al., 2000, Sullivan et al., 2006, Bartzokis et al., 2001, Courchesne et al., 2000, Yates and Juraska,

2007). Relatively little is understood about the embryonic development of the corpus callosum,

however it is known that development begins with the body of the corpus callosum known as the

genu appearing at approximately 12 weeks GA in the human and E19 in the rat (Workman et al.,

2013), later followed by the isthmus and splenium regions (Volpe et al., 2006). The developmental

gradient of the corpus callosum is unclear with some studies suggesting that is develops cranio-

caudally, while others found it to grow bi-directionally (Rakic and Yakovlev, 1968, Barkovich, 1988,

Kier and Truwit, 1997). Following birth, the corpus callosum continues to grow rapidly during

infancy, undergoing structural changes such as fibre myelination and pruning (Tanaka-Arakawa et

al., 2015, Rakic and Yakovlev, 1968). While the corpus callosum is developmentally complete by 4

years of age, it continues to expand in size at a much slower rate throughout the first 3 decades of life

(Tanaka-Arakawa et al., 2015). From 12 to 13 weeks GA, nerve fibres begin to transverse the midline

of the left and right cerebral hemispheres, laying down tracks for what will later become the corpus

callosum (Luders et al., 2010). By 18 weeks GA all regions of the corpus callosum can be visualised

(Malinger and Zakut, 1993). Myelination in the corpus callosum largely progresses in a posterior-to-

anterior direction from the splenium region to the genu (Richards, 2002, Kinney HC, 2002, Brody et

al., 1987). In the corpus callosum, the onset of myelination occurs from 9 weeks GA in the human

and postnatally from P11 in the rat (Workman et al., 2013), with the rate of myelination in humans at

its highest in childhood and although the rate declines after this, myelination persists until

approximately the 20th year of age (Keshavan et al., 2002). In rats and mice, myelinated fibres in the

corpus callosum continue development until P30 to 40, as seen using diffusion tensor imaging

(Baloch et al., 2009, Bockhorst et al., 2008). Positioned lateral and adjacent to the corpus callosum is

the external capsule, a WM tract responsible for connecting different regions of the cortex (Bloom

and Hynd, 2005). There is limited knowledge about the development of the external capsule. As the

Introduction

10

corpus callosum has a wide window of development, and is the largest WM structure in the brain, it

is susceptible to insults which impair myelination, such as IUGR.

1.4.2 Structure of the cerebellum

The cerebellum is the component of the hindbrain responsible not only for motor coordination,

balance, equilibrium and muscle tone (Buckner, 2013), but also cognition and higher order brain

functions (O'Halloran et al., 2012). The period over which the cerebellum develops extends from

early embryonic stages until the first postnatal years. This extended period of development renders

the cerebellum vulnerable to perinatal insults, which may alter cerebellar development, leading to

neurodevelopmental impairment. The fundamental mechanisms that underlie cerebellar development

in mice and rats are the same, as well as in the human, however they occur at different relative times

due to differences in gestation length. Development of the cerebellum begins with the characterisation

of the cerebellar territory at the midbrain-hindbrain boundary at E13 in the rat (Altman and Bayer,

1978). The cerebellum develops from structures known as the rhombic lips at approximately E13 in

the rat, and 5 to 8 weeks gestation in humans (Volpe, 2009). The body of the cerebellum arises from

the anterior portion of the rhombic lip, while the posterior portion, gives rise to precursors of deep

cerebellar nuclei (olivary and pontine nuclei) (Volpe, 2009). Development continues with the

formation of two compartments of cellular proliferation, the ventricular zone and the rhombic lip.

Purkinje cells and deep cerebellar nuclei arise from the ventricular zone at E13 to16 in mice, while

granule cell precursors are formed from the rhombic lip at E13 to 16 (Biran et al., 2012). Purkinje

cells are the primary output neurons of the cerebellum and aid in coordination of sensory input as

well as motor control. The Purkinje cells migrate along radial glial cells towards the pial surface and

form a mature monocellular layer at E20 to 21 (Figure 1.2), and continue to differentiate postnatally,

from P0 to P30 in the rat. The upper rhombic lip gives rise to granular precursor cells, which migrate

tangentially to form a transient germinal epithelium called the external granular layer (EGL) at E17

to 21 (Figure 1.2). Following mitotic division, the granule cells migrate, guided by Bergmann glia

(BG) to the interior of the future cerebellum, passing through the layer of Purkinje cells which are

migrating radially in the opposite direction. Purkinje cells secrete Sonic hedgehog, a protein that

stimulates proliferation of the granule precursor cells (Donkelaar et al., 2003). Once they have passed

this layer, the granule cells settle in a layer known as the internal granular layer (IGL) or granular

layer (Figure 1.2) in the mature cerebellum during the first 2 weeks after birth in humans. When

situated in the IGL, the granule cells extend horizontal parallel fibres that contact the Purkinje cell

dendrites. They are then contacted by mossy fibres, which extend in from the pons (Figure 1.2). This

developmental pattern continues, with the most rapid period of expansion occurring between 24 and

Introduction

11

40 weeks post conception in humans, resulting in a five-fold increase in the size of the cerebellum

(Chang et al., 2000).

Once development is complete, the cerebellum consists of two hemispheres, separated by a narrow

midline zone known as the vermis, and contains 3 lobes including the anterior, posterior and

flocculonodular lobe. The cerebellar anatomy consists of a tightly folded layer of GM called the

cerebellar cortex, with an underlying layer of WM, containing deep cerebellar nuclei, with the fluid-

filled fourth ventricle ventral to this. Each ridge of the cortex is known as a folium, and a set of large

folds divides the overall cerebellar structure into 10 areas known as lobules. As development of the

cerebellum is bilateral, lobules I-V arise from the anterior lobe and lobules VI to X arise from the

posterior lobe, resulting in a ‘fan shaped’ succession of lobule development from I – V and from X

to VI in each lobe. Apart from these lobules, the cerebellum is divided into many independently

functioning areas, which consist of the same stereotyped organisation of neurons, called microzones.

The outer cortex consists of four layers - the EGL which contains rapidly proliferating granular

neurons, the ML, the Purkinje cell layer and the internal granular layer IGL (Figure 1.2). The outer

molecular layer consists of the dendritic trees of Purkinje cells, Bergmann glial (BG) fibres which act

as a scaffold along which granule cells travel from the EGL to their final resting place in the IGL,

parallel fibres of granule cells which run perpendicular to the dendrites, stellate cells, basket cells and

inhibitory GABAergic interneurons which process information within the cerebellar cortex (Figure

1.2) (Volpe, 2009). The underlying Purkinje cell layer contains Purkinje cell bodies, important for

relaying information out of the cerebellum (Figure 1.2). The IGL is the deepest layer, and is densely

packed with granule cells and Golgi interneurons which filter incoming information (Figure 1.2)

(Donkelaar et al., 2003). The underlying WM is made up of myelinated nerve fibres, as well as mossy

fibres which carry information entering the cerebellum, and climbing fibres which extend from the

inferior olivary nucleus, wrap around Purkinje cells and relay information to the cells of the ML of

the cerebellar cortex (Figure 1.2).

Introduction

12

1.5 Impact of IUGR on the brain

This thesis will focus predominantly on the impact of IUGR on the largest WM tract in the cerebrum,

the corpus callosum as well as the external capsule, layer VI of the cerebral cortex and the

hippocampus. Known impacts of IUGR on these regions will be described below.

1.5.1 Impact of IUGR on the cerebrum

The cerebral cortex is the executive and integrative centre of the mammalian CNS, making up over

three quarters of the human brain’s volume (Shi et al., 2012). Cortical neurogenesis lasts over 70 days

Figure 1.2 Diagram of a mid-sagittal cross-section through the human cerebellum, with the ten lobules

indicated by roman numerals (I-X). Diagram of a mid-sagittal cross-section through the human cerebellum,

with the ten lobules indicated by Roman numerals (I-X). The dotted lines indicate the plane of section. The

cut-away illustration of the cerebellar cortical lobule depicts the three main layers of the cortex, the molecular

layer, Purkinje cell layer and granular layer. Adapted from (Ramnani, 2006).

Introduction

13

in humans (Caviness et al., 1995), and as discussed previously myelination commences in the 3rd

trimester of pregnancy and continues through childhood and adolescence into adulthood (Zhao et al.,

2005). This extended period of development renders the cerebral cortex extremely vulnerable to

perinatal insults such as inflammation/infection, hypoxic/ischaemic injury (Talos et al., 2006, Hu et

al., 2005, Hagberg et al., 2015) and IUGR induced via placental insufficiency (Rees et al., 1998, Rees

et al., 1988, Rees and Inder, 2005, Tolcos et al., 2011, Tolcos M, 2013).

A number of clinical studies using magnetic resonance imaging (MRI) to investigate the impact of

IUGR on the brain structure of infants, have reported decreased regional connectivity that is indicative

of decreased WM volume in the cerebrum, including the corpus callosum and external capsule at 12

and 18 months of age (Padilla et al., 2014b, Esteban et al., 2010), persisting through childhood and

into adulthood (Eikenes et al., 2012a). Decreased volume of the hippocampus has been reported in

IUGR infants (Lodygensky et al., 2008), as well as reduced volume of the corpus callosum (Egana-

Ugrinovic et al., 2013), and compromised folding of the cerebral cortex at post-mortem in IUGR

newborns (Dubois et al., 2008). Similarly, animal studies have shown an overall delay in myelination

of the corticospinal tract following IUGR at 52 days of gestation in guinea pigs (Nitsos and Rees,

1990), and decreased myelination in the cerebral cortex of IUGR guinea pigs at 60 and 62 dg,

persisting until 1 week of postnatal age (Tolcos et al., 2011). In the IUGR rat, reduced myelination

in the corpus callosum and external capsule is seen at P7, persisting until P14 (Azhan, A., PhD thesis,

Monash University, 2019), as well as an overall myelination delay detected at 8 weeks of age (Reid

et al., 2012) and into adulthood (Olivier et al., 2005). These studies indicate that IUGR severely

impairs cerebral development, with reduced growth of cortical and subcortical structures as well as

impaired WM development likely underlying the adverse neurodevelopmental outcomes associated

with IUGR.

1.5.2 Impact of IUGR on the cerebellum

As mentioned previously, the cerebellum increases roughly five-fold in volume between 24 and 40

weeks of GA (Volpe, 2009), and during this protracted period of growth and development, is highly

vulnerable to insults like the hypoxic-ischaemic insults occurring during IUGR. A number of clinical

studies have used MRI to investigate the impact of IUGR on the cerebellar structure of infants (Padilla

et al., 2014, Egana-Ugrinovic et al., 2013, Batalle et al., 2012, De Bie et al., 2011). Disruptions in

regional connectivity, consistent with decreased WM volume have been found in the cerebella of

IUGR infants at 1 year of age (Batalle et al., 2012), and decreased volumes of cerebellar GM and

WM have also been reported in the brains of IUGR children at 12 months, and up to 7 years of age

(De Bie et al., 2011, Padilla et al., 2011). Post-mortem studies of the IUGR brain, specifically the

Introduction

14

cerebellum are rare, however reduced TH receptor (TR) levels using immunohistochemistry (TR-α1,

α2, β1 and β2) are reduced in the cerebella of 1st and 2nd trimester IUGR foetuses (Kilby et al., 2000).

As will be discussed in more detail below (Section 1.8), TH is essential for many aspects of foetal

brain development, and thus a deficit in TR isoforms is likely to have detrimental effects on cerebellar

structure.

Animal studies examining the effect of IUGR on the cerebellum have shown reduced volumes of the

ML, IGL and cerebellar WM in the guinea pig in response to IUGR, at 60 dg, and at 1 week of age.

In the cerebellum of the IUGR guinea pig at 1 week postnatal age, neuronal density is decreased in

the IGL, as is the number of Purkinje cells (Mallard et al., 2000, Tolcos et al., 2018). An increase in

the volume of the cerebellar EGL, the site of granule cell proliferation, is seen in both the IUGR

guinea pig at 60 dg, and the IUGR rat at P7 (Tolcos et al., 2018, McDougall et al., 2017). In IUGR

compared to control rats, neuronal apoptosis is increased in the newborn (P1) cerebellum (Liu et al.,

2011), and there is a 10% reduction in the density of BG fibres at P35, which also appear disorganised

compared to controls (McDougall et al., 2017). Decreased density of BG fibres have been

documented in the cerebella of preterm human infants (Haldipur et al., 2011), and interestingly, BG

development is impaired in the presence of under-nutrition and hypoxia in rats (Benitez et al., 2014,

Clos et al., 1977). BG development appears vulnerable to IUGR, hypoxia, under-nutrition and

preterm birth, and collectively these studies display the negative impact of prenatal insults on the

developing cerebellum.

A common theme in both the IUGR cerebrum and cerebellum is reduced WM volumes. This thesis

will focus primarily on discussing and examining the decreased WM volumes, or hypomyelination

seen in the IUGR brain, and the various hypotheses speculating the mechanisms of this myelin deficit

will be discussed next.

1.6 Animal models of IUGR used to assess brain injury

A variety of IUGR animal models have been used to assess neurodevelopmental outcomes, including

those in primates (Xie et al., 2013), piglets (Bauer et al., 2004, Burke et al., 2006, Kalanjati et al.,

2017), sheep (Eixarch et al., 2012, Illa et al., 2013, van Vliet et al., 2013), guinea pigs (Lingas et al.,

1999, Mallard et al., 2000, Mallard et al., 1999, Nitsos and Rees, 1990, Tolcos and Rees, 1997, Tolcos

et al., 2011), mice (Carpentier et al., 2013, Dickinson et al., 2017) and rats (Baud et al., 2004, Caprau

et al., 2012, Ke et al., 2006, Ke et al., 2014, Lane et al., 2001b, Lin et al., 1998, Liu et al., 2011,

McDougall et al., 2017b, Olivier et al., 2007, Olivier et al., 2005, Reid et al., 2012, Tashima et al.,

2001). Animal models of IUGR have shown us that impaired WM and GM development, changes in

Introduction

15

brain volumes and neurodevelopmental and cognitive outcomes (Tashima et al., 2001, Reid et al.,

2012, Caprau et al., 2012) associated with IUGR. These models of IUGR rely on the induction of

placental insufficiency, whereby reduced blood supply to the foetus results in reduced oxygen,

nutrients and hormones for development. Placental insufficiency is initiated via uterine artery ligation

(one or both arteries; from E17 to 19) in rats (Term = E22), rabbits (~25 days of gestation; Term =

31 days), and single umbilical artery ligation in sheep. This thesis will focus on an established rat

model of IUGR. There are a number of reasons this model of IUGR was chosen for our study.

Importantly, key time-points of brain development and in particular myelination and OL development

have been well characterised in the rat, including the Wistar strain (Craig et al., 2003). At the time of

BUVL in our model of IUGR (E18), preOLs have already begun to colonise the brain (Richardson et

al., 2006). OL maturation and myelination occurs mostly after birth in the rat from P1 to 14, and this

timeframe is equivalent to the development of myelin in the human brain at 23 to 40 weeks of GA

(Craig et al., 2003, Semple et al., 2013)(Figure 1.5). In the rat, the development of myelin in the brain

at P2 to P5 is similar to the human brain at 23 to 32 weeks GA (Segovia et al., 2008, Wu et al., 2013),

and at P7 cerebral myelination is histologically similar to WM in the human brain at 32 to 34 weeks

GA (term = 40 weeks) (Vannucci and Vannucci, 1997). This model also mimics the clinical effects

of IUGR, including reduced body (Lane et al., 2001b, Olivier et al., 2005, Price et al., 1992, Ogata et

al., 1985, Sadiq et al., 1999) and brain (Sadiq et al., 1999, Ogata et al., 1985, Tashima et al., 2001)

weight.

1.7 Mechanisms of reduced and delayed myelination in IUGR

Myelin is a lipid-rich substance containing proteins, and is essential for insulating nerve axons in the

brain, allowing for electrical impulses to be propagated rapidly and efficiently to enable correct motor

and cognitive function. In humans myelination begins in the second half of gestation and peaks during

the first year of life (Jakovcevski et al., 2009), continuing into adulthood (Richardson et al., 2011).

In the CNS, myelin is produced by OLs, which undergo a lineage progression. At least four stages of

OL maturation have been characterised and include OL progenitor cells (OPCs), pre-

oligodendrocytes (pre-OLs), pre-myelinating OLs and mature myelinating OLs (Figure 1.3).

Myelination is disrupted in the IUGR brain, and several underlying mechanisms have been proposed,

including decreased blood flow (or ischaemia) to WM regions causing a reduction in myelin

production, thought to be due to impaired protein function and plaque-like aggregates (Zhan et al.,

2015); WM regions are thought to be more vulnerable to an ischaemic insult than GM due to more

sparse capillaries (Volpe, 2001, Greisen and Borch, 2001). An alternate hypothesis is that IUGR

delays myelination through the generation of oxidative stress, whereby reactive oxygen species,

thought to be produced by mitochondria in the presence of hypoxia-ischaemia (Kirkinezos and

Introduction

16

Moraes, 2001), reduce the expression of genes that promote OL lineage maturation, and elevate those

which inhibit maturation (French et al., 2009). Supporting this hypothesis, Reid and colleagues

showed that IUGR in the rat generates reactive oxygen species postnatally, which in turn up-regulates

bone morphogenic protein 4 (Reid et al., 2012), a known inhibitor of OL differentiation. There is

substantial evidence for the susceptibility of the OL lineage to oxidative stress, however OL

progenitors as well as pre-myelinating OLs are more susceptible than mature OLs, as they containing

lower levels of antioxidants (Back et al., 2002). It has been proposed that developmental window in

which the brain contains the highest numbers of immature OLs is the time at which it is most

vulnerable to insults like those occurring as a result of IUGR. This timeframe of vulnerability

translates to < 30 weeks GA in humans (Back et al., 2001), and from P7 to P10 in rats (Craig et al.,

2003).

Additionally in the cerebral cortex of IUGR foetal guinea pigs compared to controls, an abnormal

retention of myelin basic protein (MBP) in the OL cell body, and lack of MBP-IR staining in the OL

processes, has been reported (Tolcos et al., 2011). MBP mRNA is normally translocated from the OL

soma into its distal processes, and these processes ensheath neuronal axons during the myelination

process (Pedraza, 1997), thus this result in the IUGR foetal guinea suggests a delay in the production

and/or trafficking of MBP proteins necessary for axonal myelination. While IUGR may delay the

migration of key myelin proteins, recent studies in guinea pigs and rats have explored the hypothesis

that IUGR delays the maturation of OLs, halting them at the pre-myelinating stage, therefore resulting

in delayed myelination in the brain (Reid et al., 2012, Tolcos et al., 2011, Rideau Batista Novais et

al., 2016), which persists after birth (Olivier et al., 2005). A study by our group using a rat model of

IUGR (bilateral uterine vessel ligation (BUVL) in late gestation; E18) revealed an increased

expression of genes that inhibit OL maturation in the IUGR brain including Axin2, Notch1 and 2,

Jagged1, Hes5 and BMP4, and a decrease in genes that promote OL maturation, such as Sox10 and

Myrf (Azhan, A., PhD thesis, Monash University, 2019). Impaired myelination in IUGR is likely to

occur due, in part, to all of these discussed mechanisms. Another possible mechanism, which is being

explored and is relevant to the current study, is impaired cerebral TH signalling.

Introduction

17

TH is essential for brain development, including for the maturation of the OL lineage as well as for

the process of myelination (Barres et al., 1994). TH signalling will be discussed in greater detail

below (Section 1.7.1), however it is important to note that TH enters cells via intercellular

transporters, with MCT8, alternately known as ‘solute carrier family 16, member 2’ (SLC16A2),

exclusive to transportation of TH (Friesema et al., 2003). Of interest, MCT8 levels are reduced in

IUGR. A post-mortem study found that MCT8 expression was significantly reduced in the cortical

plate of foetuses with severe IUGR compared to non-IUGR foetuses (Chan et al., 2014). Reduced

mRNA levels of MCT8 are also shown in the brain of IUGR rats at P7 (Azhan, A., PhD thesis,

Monash University, 2019). Interestingly in clinical cases of MCT8 mutation, persisting deficits in

myelination are seen in the brain. A post-mortem study examining brain tissue from a foetus, and an

11 year old male with MCT8 mutations, showed significant myelin deficits in the cerebral cortex as

well as a 50% reduction of TH (T3, 3,5,3’-triiodothyronine; and T4, thyroxine) (Lopez-Espindola et

al., 2014). In children with Allan-Hernon-Dudley syndrome (AHDS), characterised by an MCT8

mutation, significant and lasting hypomyelination is also seen in the brain (Verge et al., 2012).

Together these studies highlight the importance of MCT8 and cerebral TH signalling in the process

Figure 1.3 Timeline of oligodendrocyte development from pre-oligodendrocytes to the mature

myelinating phenotype in humans and rats. Pre-oligodendrocytes (A; pink bars) developing into immature

non-myelinating oligodendrocytes (B; blue bars) and then into a mature myelinating phenotype (C; orange

bars). This lineage maturation occurs postnatally in the rat, but commences prior to birth in the human,

continuing into adulthood. Mature oligodendrocytes lay down myelin insulating neuronal axons in the brain.

Figure adapted from Semple et al. 2013 and Jakovcevski et al. 2009.

Introduction

18

of myelination, and suggest that deficits in MCT8 seen in IUGR, along with impaired TH signalling

may underpin the mechanism behind disrupted WM and myelin development.

1.8 Impact of IUGR on thyroid gland function

TH is made by the thyroid gland, and is essential for growth and development. Before the effects of

IUGR on thyroid gland function and circulating TH levels is discussed further, a brief overview of

thyroid gland function, as well as the cellular TH signalling pathway will be provided.

1.8.1 Overview of TH signaling

Anatomically, the thyroid gland is situated between the 5th cervical and 1st thoracic vertebrae, ventral

of the trachea and below the larynx, and it consists of 2 lobes which are connected by an isthmus

(Stathatos, 2006). The thyroid gland produces and releases TH into the circulation. The brain, heart,

lungs, kidneys, liver, intestines and reproductive organs are all reliant on TH signalling for

development and proper function (Peeters et al., 2003, Warner and Mittag, 2012). The thyroid gland

uses iodine, absorbed through the gastrointestinal tract to synthesise THs, with the predominant forms

of TH including 3, 5, 3’, 5’- tetraiodo-L-thyronine, or thyroxine (T4), and triiodothyronine (T3)

(Figure 1.4). A large percentage of THs bind to TH distributor proteins (THDPs) in the plasma

(Schreiber and Richardson, 1997), with known THDPs including albumin, thyroxine-binding

globulin, transthyretin and certain lipoproteins (Prapunpoj et al., 2000, Lans et al., 1994, McLean et

al., 2017). Less than 1% of circulating THs remain unbound and are known as ‘free’ T3 and T4 (FT3,

FT4), and this small fraction is biologically active (Mendel, 1989). T3 is 5 times more biologically

active than T4, due to being bound less tightly to transporter proteins, and as a result is more readily

available for cellular binding and uptake (Schroeder and Privalsky, 2014).

THs enter cells using various transmembrane transporters (Figure 1.4), including anion transporting

polypeptides, L-type amino acid transporters and monocarboxylate transporters. Of particular

importance to this thesis, MCT8 is the only transporter to exclusively transport THs in the brain (Di

Cosmo et al., 2009). Once inside the cell, T4 undergoes a process called deiodination, catalysed by

iodothyronine deiodinases (Type-1; Dio1, Type-2; Dio2, Type-3; Dio 3) (Gereben et al., 2008) to

remove iodine atoms from the outer ring of T4 (Visser et al., 1988), producing active T3. T3 can then

enter the nucleus and bind to nuclear TRα and β (Evans, 1988) and regulate the expression of specific

genes in that cell. TH signalling in the body relies on a negative feedback mechanism to maintain

homeostasis. For example, if circulating TH levels in the blood become too low, the hypothalamus

secretes thyrotropin-releasing hormone (TRH) which stimulates the pituitary gland to secrete thyroid-

Introduction

19

stimulating hormone (TSH), signalling the thyroid gland to release more T3 and T4 into the blood

stream, thus normalising circulating TH levels. Conversely, if circulating TH levels are too high, the

hypothalamus down-regulates its secretion of TRH, ultimately leading to the thyroid gland releasing

less T3 and T4 into circulation (Maria Izabel Chiamolera, 2009). In IUGR foetuses, the literature

surrounding circulating TH levels is inconclusive, with studies finding reductions in T4 and FT4

levels yet no difference in T3 or FT3 compared to non-IUGR foetuses (Thorpe-Beeston et al., 1991,

Soothill et al., 1992). Others have found that both T3 and T4 are reduced in IUGR foetuses compared

to non-IUGR foetuses (Kilby et al., 1998), and that in IUGR infants, plasma T3 and T4 levels are

reduced at birth (Bongers-Schokking and Schopman, 1984, Jacobsen and Hummer, 1979). T3 levels

however, begin to normalise from 1 week of postnatal age until 8 months of age, and T4 begins to

normalise from 2 months of age (Jacobsen and Hummer, 1979). Animal studies examining TH levels

in offspring following IUGR are rare, however a study of growth restricted lambs (induced via

carunclectomy prior to pregnancy conception) reported decreased plasma T4, and increased T3 levels

postnatally (De Blasio et al., 2006). These conflicting results make it difficult to determine the TH

profile of IUGR infants, thus further studies examining postnatal TH levels are required, especially

considering the essential role of TH in brain development.

Introduction

20

1.9 Role of TH in brain development

In the brain, TH is essential for driving neurogenesis, synaptogenesis, and cell migration (Younes-

Rapozo et al., 2006, Barres et al., 1994, Berbel et al., 1994, Berbel et al., 2010, Skeaff, 2011, Zoeller

and Rovet, 2004), while acting as a fundamental driver of OL maturation and differentiation as well

as the expression of myelin specific genes (Reid et al., 2012). The foetus relies on maternal TH levels

until approximately 20 weeks GA, after which time the foetus’s thyroid gland begins releasing its

own hormones, causing foetal TH levels to rise during late gestation, peaking around the time of birth

(Figure 1.5 A). During this time the brain undergoes major developmental changes, including

development of the neocortex and cerebellum, glial cell proliferation, synaptogenesis, axonal and

dendrite sprouting and neuronal migration and proliferation (Figure 1.5 B). For the purposes of this

thesis, The role of TH in the development of myelin within the brain will be a focus. In rats, a lack of

TH (hypothyroidism) results in decreased myelination in the foetal brain (Calvo et al., 1990),

persisting during the 1st postnatal month (Rodriguez-Pena et al., 1993), while an excess of TH

(hyperthyroidism) results in a significant increase in myelination at P13 (Walters and Morell, 1981).

T

Thyroid gland

T4

Circulation

T4

T4

T4

T4

Lat1

MCT10

OATP1C1

MCT8

T3

Dio1 or Dio2

T2 rT3

Dio3 Dio3

T4

Target cell

TRE

RXR

Myelin RNA

Myelin Protein

T3

T3

Dio1 or Dio2

Nucleus

TR α/β

Figure 1.4 Thyroid hormone cellular signalling pathway. This schematic uses the example of thyroid

hormone (TH; T3 and T4) entering an oligodendrocyte to promote the transcription of myelin protein. Thyroid

hormone is released from the thyroid gland and circulates around the body via the blood. It contacts and enters

cells using transporters, with MCT8 being exclusive to TH transport. Once inside the target cell T4 is

deiodinated into an active T3 form by various deiodinase enzymes (Dio1 & Dio2), and enters the nucleus,

binding to TH receptors alpha and beta (TRα, TRβ) where it promotes the expression of mRNA and proteins.

Dio1 = deiodinase 1, Dio2 = deiodinase 2, Lat1 = large amino acid transporter 1 MCT10 = monocarboxylate

transporter 10, MCT8 = monocarboxylate transporter 8, OATP1C1 = Organic anion transporting polypeptide,

RNA = Ribonucleic acid, RXR = retinoid X receptor, T3 = triiodothyronine, T4 = thyroxine, TR = thyroid

hormone receptor. Original figure.

Introduction

21

Given that myelin is essential for the efficient and correct propagation of action potentials in the brain,

it is possible that the poor cognitive outcomes associated with IUGR may be due impaired

myelination caused by deficits in cerebral TH signalling. Indeed neurodevelopmental impairment

occurs in disorders of hypothyroidism, including when TH is deficient in the maternal circulation

during gestation, and in congenital hypothyroidism which typically occurs as a result of dysgenesis,

or malfunction of foetus’s thyroid gland. Neurodevelopmental impairments resulting in reduced

intelligence quotient scores (IQ) and speech delays and deficits have been implicated in both types of

hypothyroidism (Selva et al., 2005, Haddow et al., 1999). Furthermore autism (Román et al., 2013)

and attention deficit hyperactivity disorder (ADHD) (Vermiglio et al., 2004), two disorders with

known neurodevelopmental impairment, are associated with TH deprivation in utero. Disorders of

TH signalling such as congenital hypothyroidism highlight the important link between TH and correct

brain development.

Introduction

22

Several categories of hypothyroidism exist, including congenital hypothyroidism (CH) which is

characterised by being present from the time of birth (LaFranchi, 1999), autoimmune hypothyroidism

such as Hashimoto’s thyroiditis in which inflammation (lymphocytic thyroiditis) renders the thyroid

gland underactive (Lorini et al., 2003), hypothyroidism arising due to iodine deficiency (Patrick,

2008), pregnancy (Kothari and Girling, 2008), or due to medical intervention including surgery or

chemotherapy (Bettendorf et al., 1997, Beltran et al., 2006, Desai et al., 2006). This section will focus

on CH, which is characterised by deficient circulating TH levels at the time of birth, and is of

particular relevance to this thesis, as it causes neurodeficits which are similar to those seen in IUGR,

including hypomyelination.

Figure 1.5 Timing of human and rat brain development in relation to thyroid hormone signalling. (A)

Thyroid gland development and TH gradients throughout gestation and early childhood. (B) Key

developmental time points for brain development. Development of the thyroid gland occurs between 3 and

13 weeks GA in the human, and foetal TH secretion peaks at birth in the human, and postnatally in the rat.

(B) Brain development, including myelination in humans has largely commenced prior to birth and

continues into childhood and adolescence. In the rat, myelination begins after birth, as does glial cell

proliferation, synapse formation and axonal spouting. Yellow bar = time frame for term birth in rats. Blue

dotted lines indicate when process is active, and solid blue bars indicate the dominant window of time in

which the process occurs. D2 = deiodinase 2, TH = thyroid hormone, TSH = thyroid stimulating hormone,

T3 = triiodothyronine. Image adapted from Bernal et al. 2007; Lemkine et al. 2005; Remaud et al. 2017.

Introduction

23

CH is defined by partial or full loss of thyroid gland function during foetal development, and is

divided into primary, secondary and peripheral etiologies. Primary CH occurs due to either dysgenesis

of the thyroid gland (~85% of cases) (Rastogi and LaFranchi, 2010), or direct impairment in TH

biosynthesis (~10 to 15% of cases) (Rastogi and LaFranchi, 2010). Secondary, or central

hypothyroidism is associated with pituitary hormone dysfunction leading to TSH deficiency and

decreased circulating T3 and T4 levels (Rastogi and LaFranchi, 2010), while peripheral CH is

identified by impaired TH transport, metabolism or action (Misiunas et al., 1995). CH can be transient

or permanent, with permanent CH persisting throughout life and requiring ongoing treatment, while

transient CH is discovered at birth but can be alleviated with treatment in the first few years of life

(Razavi and Mohammadi, 2016).

Various clinical studies have demonstrated neurodevelopmental impairments in children in cases of

both transient and permanent CH, with these impairments including deficits in cognitive

development, language and speech (Gottschalk et al., 1994), visuospatial processing (Leneman et al.,

2001), attention (Kooistra et al., 1996), motor skills (Cooper et al., 2019, Fuggle et al., 1991), and

auditory function (Peters et al., 2018). A number of trials have reported IQ scores as a measurement

of the impact of CH on neurodevelopment, and have correlated these IQ outcomes with the timing at

which TH replacement therapy commenced (Hindmarsh, 2002, Bongers-Schokking et al., 2000,

Selva et al., 2005, Selva et al., 2002, LaFranchi, 1999, Dubuis et al., 1996, Rovet and Ehrlich, 1995,

Simoneau-Roy et al., 2004. Children with either transient or permanent CH have IQ scores that are

on average 6 points below their euthyroid counterparts (Najmi et al., 2013, Ordooei et al., 2014,

Derksen-Lubsen and Verkerk, 1996). Infants with CH are typically treated with Levothyroxine (L-

thyroxine), a synthetic form of TH which behaves identically to endemic T4. Countless studies show

early intervention to be essential in CH, with infants who commence TH replacement therapy during

the neonatal period displaying superior neurodevelopmental outcomes when assessed during

childhood compared to those treated later on (Hindmarsh, 2002, Bongers-Schokking et al., 2000,

Selva et al., 2005, Selva et al., 2002, LaFranchi, 1999, Dubuis et al., 1996, Rovet and Ehrlich, 1995,

Simoneau-Roy et al., 2004). Animal studies also show the advantage of early intervention. In

hypothyroid rat pups (induced via maternal consumption of Methimazole; 7.5µg/day in drinking

water; E16 – P25) treated with T4 replacement therapy (0.02µg/g body weight; from P1 – P21),

improvements in behavioural testing scores such as perseverance, memory and learning were reported

only following the first 8 days of treatment (from P1-P7), with no improvements beyond this time

(MacNabb et al., 2000, O'Hare et al., 2015, Reid et al., 2007).

Introduction

24

In the brain, CH impairs myelination in neonates, but this can be reversed with TH replacement (T4)

therapy (Jagannathan et al., 1998), and delayed myelination is evident at 1 year of age using MRI

(Ari Yuca 2014), while altered microstructure of WM is reported in childhood, shown by decreased

fractional anisotropy in the cerebellum, thalamus and temporal lobe using diffusion-MRI (Cooper

2019). In children (6 to 15 years of age), changes in metabolites which are indicative of myelination

arrest are observed using MRI (Gupta et al., 1995); these outcomes were also reversed using T4

replacement therapy during childhood (Gupta et al., 1995). Animal studies similarly show impaired

neurodevelopment as a result of CH, including impaired myelination. Post-mortem analysis in dogs

with CH, revealed myelin deficits in the corpus callosum and pons (Pettigrew et al., 2007), while in

rats, CH reduced myelination in the anterior commissure region of the brain (Lucia et al., 2018), and

decreased mRNA levels of MBP and PLP (Rodriguez-Pena et al., 1993). Reduced myelin deposition

is correlated with impaired axonal maturation in the brains of CH mice (Noguchi and Sugisaki, 1984)

and dogs (Pettigrew et al., 2007), suggesting that the hypomyelination seen in CH is a consequence

of impaired axonal formation. Collectively, these studies highlight the fundamental importance of TH

in brain development, the implications of abnormally decreased TH levels for myelination in the

brain, and the importance of treating these abnormal levels as early as possible.

1.10 Use of thyroid hormone in neonatal brain injury

Varied results have been reported in human and animal studies examining the use of TH, and TH

analogues in the treatment of neonatal brain injury. In a study of newborns with congenital

hypothyroidism treatment with T4 was able to normalise myelin structure within the cerebellum and

cortical WM, reversing abnormal lipid peaks when assessed using magnetic resonance spectroscopy

(Jagannathan et al., 1998). In contrast, a study examining the neurodevelopmental benefits of T4

treatment in very preterm infants (born 26 to 28 weeks GA), found no benefit for the impaired

neurodevelopmental outcomes associated with very preterm birth including reduced IQ and

developmental delay (Chowdhry et al., 1984, Lucas et al., 1996, Meijer et al., 1992, van Wassenaer

et al., 1997, Vanhole et al., 1997). Animal models show benefits of TH treatment for neonatal brain

injury, with no adverse effects. In a rabbit model of intraventricular haemorrhage, treatment with T4

(20µg/day) from 24 hours to 10 days postnatally restored myelination within the brain (Vose et al.,

2013). Similarly, in rat (Nathaniel et al., 1988) and mouse (Noguchi et al., 1985) models of

hypothyroidism, treatment with T4 (2.5µg/day commencing P14 or P20 until P42)(Nathaniel et al.,

1988) or 0.1µg T4 daily injections from birth until P20 (Noguchi et al., 1985) improved brain weight

and a reversed of cerebral hypomyelination. Improvements in the mouse model of hypothyroidism

were only seen when T4 was given within the first 20 postnatal days (Noguchi et al., 1985), with P20

in rodents equivalent to approximately 1 year of age in humans (Porterfield and Hendrich, 1993). The

Introduction

25

results of these studies indicate that TH replacement therapy may be beneficial for restoring normal

levels of myelination in the brain when given to hypothyroid subjects, with emphasis placed on

commencing treatment in the neonatal period immediately after birth. When brain injury is induced

by inflammation however, TH replacement therapy may not be effective. In a neonatal mouse model

of systemic inflammation (induced by IL-1β), treatment with T4 (20µg/kg/day from P1 to P5) was

unable to reverse the block in OL maturation within the brain caused by inflammation when assessed

at P1 and P30 (Schang et al., 2014).

TH analogues, including levothyroxine (LT3), 3,3’,5-triiodothyroacetic acid (Triac), 3,3’,5,5’-

tetraiodothyroacetic acid (Tetrac) and 3,5-diiodothyropropionic acid (DITPA) have also shown

varied potential as therapies for neonatal brain injury; the use of DITPA will be discussed in greater

detail below (Section 1.10). A multicentre clinical trial of extremely preterm infants given LT4

(8µg/kg birth weight/day; within 5 days of birth until 32 weeks corrected GA), measured the

therapeutic benefits of LT3 therapy in reducing neurodevelopmental disability associated with

hypothyroidism in preterm birth, but found no improvement in disability outcomes (Ng et al., 2014).

In animal models, treatment with TH analogues Triac and Tetrac have varied results. In mice deficient

in the exclusive TH transporter MCT8, Tetrac (400ng/g body weight; daily; first 2 postnatal weeks)

promoted TH dependent gene expression in the brain (Horn et al., 2013). Triac (400ng/g body weight;

daily injection) similarly prevented neuronal damage in the brains of newborn mice who were

hypothyroid (Kersseboom et al., 2014). In contrast, when administered directly into the brain of

MCT8 deficient mice, Triac failed to replace TH in mediating the actions of target genes within the

brain (Barez-Lopez et al., 2019).

TH analogue therapies have also been explored in the treatment of AHDS. This disorder is

characterised by an MCT8 mutation, thought to delay TH transport into cells in the brain, causing

delayed myelination with intellectual and physical disability (Schwartz and Stevenson, 2007).

Treatment of 4 children with AHDS on compassionate grounds with DITPA (1.8mg initially; 30mg/d

[2.1 – 2.4 mg/kg/day] given in three divided doses, orally 26 – 40 months), which enters cells

irrespective of MCT8, improved myelination in the brains of 2 out of 4 children when assessed using

MRI (Verge et al., 2012). Treatment with conventional TH in IUGR is likely to be ineffective, as

expression of the exclusive TH transporter MCT8 is reduced in IUGR (Azhan, 2019, Chan et al.,

2014), therefore alternate therapies including TH analogues such as DITPA, which do not rely on

MCT8 for cellular uptake, should be investigated.

Introduction

26

1.11 DITPA as a therapy in adults and children

DITPA is a synthetic TH analogue which has similar biological properties to endogenous TH. DITPA

acts as a TR agonist, binding to both and isoforms in the cell nucleus (See Figure 1.4 for TH

signalling) (Pennock et al., 1992, Raparti et al., 2013), and a similar structure to T3 but lacking one

iodine molecule and an NH2 group (Figure 1.6). As previously mentioned, DITPA has been

investigated as a therapy for the hypermetabolism and weight loss associated with elevated circulating

TH levels in AHDS (Verge et al., 2012), as it does not require MCT8 to be transported into the cell

(Di Cosmo et al., 2009). There is also evidence that DITPA can cross the blood-brain barrier, as well

as the placenta (Ferrara et al., 2015). In a case report of 4 children with AHDS, DITPA treatment

(1.8mg initially; 30mg/d [2.1 – 2.4 mg/kg/day] given in three divided doses, orally 26 – 40 months)

was well tolerated without thyrotoxic effects, and normalised thyroid function and heart rate as well

as improved weight gain (Verge et al., 2012). With human (Chan et al., 2014) and rodent (Azhan, A.,

PhD thesis, Monash University, 2019) data to indicate that MCT8 is reduced in the IUGR brain, the

study by Verge and colleagues (Verge et al., 2012) provides reason to think DITPA would be

beneficial when given in cases of IUGR also. Clinical trials in adults with cardiac failure similarly

found that DITPA treatment did not cause hypothyroidism or thyrotoxicity (Goldman et al., 2009,

Ladenson et al., 2010), although it did cause gastrointestinal upset, likely due to the large dose given

(90 to 360 mg/day). In MCT8 knockout mice, DITPA (0.3mg/100g body weight/day) ameliorated a

thyrotoxic liver state and normalised plasma T3 to control levels (Di Cosmo et al., 2009). These

studies support the safety and benefits of DITPA, however little is known of the benefits of DITPA

therapy when given to IUGR infants. Our group has previously investigated the efficacy of DITPA

treatment (0.5mg/100g i.p. daily), given to IUGR rat pups from P1 to P6, and by P7 found it was well

tolerated, did not cause adverse effects on body weight, body composition or brain weight (Azhan,

A., PhD thesis, Monash University, 2019).

Introduction

27

1.11.1 Physiological impacts of DITPA treatment.

The clinical safety profile of DITPA has been reported in children with mutations in the MCT8 gene

(Verge et al., 2012), adults with cardiovascular disease (Goldman et al., 2009), and in MCT8

knockout mice (Di Cosmo et al., 2009). The results of these important human and animal studies are

listed in Table 1.1 below, outlining DITPA’s impact on various parameters such as growth, body

weight, body composition, thyroid function, liver function and cholesterol levels. The broad

consensus from these studies is that DITPA treatment did not cause harm, except for reducing body

weight in adults with cardiac failure and increasing markers of bone turnover (Goldman et al., 2009,

Ladenson et al., 2010) when given as a very large dose (90 – 360mg/d; orally) over a relatively short

duration of time (8 or 24 weeks). In children with AHDS (x-linked MCT8 mutation; gene locus

SLC16A2) given a smaller dose (30mg/d; orally) for a longer duration of time (26 – 40 months),

DITPA improved weight gain and normalised circulating TH levels (Verge et al., 2012). In MCT8

knockout mice, DITPA once again normalised TH levels and ameliorated a thyrotoxic liver state (Di

Cosmo et al., 2009). When DITPA was administered to IUGR rat pups (0.5mg/100g/day i.p injection

Figure 1.6 Chemical structure of thyroid hormones and analogues: (A) Triiodothyronine; T3, (B)

thyroxine; T4 and (C) 3, 5-diiodothyropropionic acid (DITPA). DITPA is structurally identical to T3 and T4

albeit for the lack of an amino radical group (NH2; yellow circle shows where absent), and differs from T3 by

the absence of an iodine atom (blue circle), and from T4 by two iodine atoms (blue and green circles). I =

iodine, NH2 = amino radical. Figure adapted from Davis et al. 2009.

Introduction

28

from P1 to P6), brain-to-body-weight ratio was improved compared to pups given saline, which

indicated improved brain growth, however DITPA decreased lean tissue mass, fat mass and bone

mineral density in IUGR pups at P7 (Azhan, A., PhD thesis, Monash University, 2019), suggestive

of increased bone turnover like that seen in humans (Table 1.1).

Introduction

29

Reference Patients/animal

model

DITPA regimen –dose,

route and duration

Type of

control

Impact of DITPA

on growth & body

weight

Impact of

DITPA on body

composition

Impact of DITPA on

thyroid and liver

function

Goldman et al.

2009

86 adults with

congestive heart

failure (> 18yrs).

Twice daily, oral dose

increased in 90mg/d

increments every 2 weeks

to maximum 360mg/d for

24 weeks.

Stage II multicentre

randomised, double-

blinded trial.

Placebo

capsules. • Decreased body

weight (-11lb).

Not assessed • Decreased plasma

cholesterol (- 20 %.)

• TSH was supressed.

Di Cosmo et al.

2009

Adult male mice

with MCT8 gene

mutation (MCT8-

knockout).

03, 0.6 or 1.0mg/100g body

weight/day, i.p. injection for

4 days.

Wild type

male mice

(no

MCT8-

knockout)

• Not assessed Not assessed • Higher dose

normalised plasma

TSH.

• Ameliorated

thyrotoxic liver

state.

Ladenson et al.

2010

86 adults with

chronic heart

failure (55 to 77

yrs).

Escalating oral dose,

90mg/d to 180mg/d,

270mg/d, and 360 mg/d

over 8 weeks.

• Pilot prospective,

randomized controlled

study.

Placebo

capsules. • Decreased body

weight.

Indication of

increased bone

turnover.

• Lowered plasma

TSH levels.

• Lowered plasma T3

& T4.

• No hypothyroidism.

• No thyrotoxicosis.

• Decreased plasma

cholesterol.

Introduction

30

Table 1.1 Important DITPA studies in humans and animals. (Table continued on next page).

d = day, g = grams, i.p. = intraperitoneal, IUGR = intrauterine growth restriction, kg = kilograms, lb. = pounds, MCT8 = monocarboxylate transporter – 8, mg

= milligrams, P = postnatal day, T3 = triiodothyronine, T4 = thyroxine, TSH = thyroid stimulating hormone, yrs = years of age.

Verge et al. 2012 4 children with

MCT8 mutation,

treatment

commencing 8.5

to 25 months of

age.

1.8mg initially; 30mg/d

(2.1 – 2.4 mg/kg/day)

given in three divided

doses.

Oral for 26 to 40 months.

• DITPA given on

compassionate grounds.

None. • Improved weight

gain in 2/4

children.

Not assessed • Normalised TH

levels (normalised

elevated T3 & TSH,

increased T4).

Ferrara et al. 2015 Adult male mice

with MCT8 gene

mutation (MCT8-

knockout).

0.3 mg/100g body

weight/day, i.p. injection,

for 10 days, commencing

P7.

Wild type

male mice

(no

MCT8-

knockout)

Not assessed Not assessed • Normalised plasma

TSH and T3.

• Ameliorated

hypermetabolism.

Azhan 2019 PhD

thesis, Monash

University, 2019.

Newborn IUGR

rats (IUGR

induced by

bilateral uterine

vessel ligation in

late gestation).

0.5mg/100g/day i.p injection

from P1 to P6 (i.e. 7 days).

Saline i.p.

injection.

In IUGR pups at P7:

• Increased brain-to-

body-weight ratio.

In IUGR pups at

P7:

• Decreased lean

tissue mass.

• Decreased fat

mass.

• Decrease in

bone mineral

density

Not assessed

Introduction

31

1.11.2 Impact of DITPA on the CNS and developing brain

The effects of DITPA on the developing brain are largely unknown, as are the mechanisms by which

DITPA enters cells in the brain. It is known however, that DITPA (1mg/100g/day) readily enters

brain cells in MCT8 knockout mice, and subsequently corrects cerebral hypothyroidism (Di Cosmo

et al., 2009). A previous study by our group found that DITPA (0.5mg/100g; i.p.) administered daily

from P1 to P6 in IUGR rat pups, promoted myelin recovery, detectable by P7 in the external capsule

(Azhan, A., PhD thesis, Monash University, 2019). In cases where MCT8 transporters are deficient,

such as in IUGR, the ability of endogenous TH to enter cells such as OLs and initiate the transcription

of myelin specific genes is impaired. This contributes to the delay in OL development and

myelination seen. DITPA bypasses MCT8 and may act to replace endogenous TH, thus promoting

OL development and myelination, although the exact mechanisms by which DITPA acts on genes

associated with myelination is unknown. Although promising, this data only provides evidence of the

impact of short-term DITPA treatment on the cerebrum, and although benefits were seen in the

external capsule, a longer dosing regimen may further benefit the brain. There is currently no

treatment to prevent IUGR. If detected in utero, the only intervention is to deliver the baby preterm

to remove it from the compromised intrauterine environment dominated by the insufficient placenta;

this typically occurs any time between 24 and 40 weeks GA. Once removed from the unfavourable

intrauterine environment, DITPA therapy could be given to an IUGR baby soon after birth; however

the dose, duration and route still need to be determined. Before such clinical studies are undertaken,

it is first important to test the preclinical efficacy and safety of DITPA in a preclinical model of IUGR.

1.12 Scope of this thesis

It is proposed that the synthetic TH analogue DITPA, which does not require MCT8 to enter cells,

can be used to overcome the deficits caused by a loss of MCT8 expression in the IUGR brain and

restore TH uptake, thereby normalising myelination and OL development. Taking into account the

findings of our group’s previous study whereby MCT8 levels normalised by P14 in the IUGR rat,

this project set out to investigate the benefits of DITPA administration from P1 to P13 a time in rat

brain development equivalent to that of a human baby at 23 to 40 weeks GA (Semple et al., 2013),

and reflective of what is likely to occur in the clinical setting with an IUGR baby delivered preterm

and given DITPA until term equivalent age. If serious, IUGR is detected during late pregnancy, and

the baby is often delivered preterm in order to remove it from the insufficient intrauterine

environment. The longer-term DITPA administration used in this study (P1-P13) importantly

addresses the clinical scenario of an IUGR baby being delivered preterm, and then being administered

DITPA until term equivalent age. This model of IUGR was ideal, as OL maturation and myelination

occurs postnatally in the rat from P1 to 14, and this timeframe is equivalent to the development of

Introduction

32

myelin in the human brain at 23 to 40 weeks of GA (Craig et al., 2003, Semple et al., 2013). In the

rat, the development of myelin in the brain at P2 to P5 is similar to the human brain at 23 to 32 weeks

GA (Segovia et al., 2008, Wu et al., 2013), and at P7 cerebral myelination is histologically similar to

WM in the human brain at 32 to 34 weeks GA.

The overall aim of this thesis was to determine the potential of DITPA administration

(0.5mg/100g/day i.p. from P1 to P13) to restore myelination in the IUGR brain and promote OL

development, without causing brain injury or neuroinflammation, or any negative off-target effects

on the body. Below, the aims and objectives of each of the studies, which make up the experimental

Chapters in this thesis will be outlined.

Chapter 3: In Chapter 3, the impact of DITPA administration on the development of the cerebrum

was investigated, focusing on the cortical layer VI, corpus callosum, external capsule, hippocampus

and fimbria as these regions are known to be vulnerable to injury in IUGR (Padilla et al., 2011, Padilla

et al., 2014b, Egana-Ugrinovic et al., 2014, Lodygensky et al., 2008). Assessment of myelination has

been a focus, as this is one of the main aspects of brain development that is affected in IUGR the

human (Chase et al., 1972, Eikenes et al., 2012a, Esteban et al., 2010, Padilla et al., 2014b) and rat

(Azhan, 2019, Olivier et al., 2005, Reid et al., 2012). This study aimed to determine whether longer-

term DITPA treatment in IUGR rat pups (from P1 to 13) (i) restored myelination, (ii) promoted OL

maturation, and (ii) did not cause injury or inflammation, in the cerebrum. It was hypothesised that

DITPA would improve myelination and OL development in the cerebrum of IUGR pups, without

causing injury or inflammation. The results of this study suggest that DITPA treatment in IUGR rat

pups, promotes immunoreactivity (IR) for MBP in cortical layer VI and the fimbria, and Olig2-IR

OL density in the corpus callosum when assessed at P14, but reduces the length of MBP-IR

projections in the cortex, and may impair expression of myelin protein PLP-IR in the corpus callosum

and external capsule. DITPA does not cause injury or inflammation in any of the cerebral structures

investigated, however leads to unfavourable outcomes when given to control pups, including

decreased PLP-IR in the corpus callosum and external capsule, and decreased density of Olig2- and

APC-IR OLs in cortical layer VI and fimbria. With indication that this duration of DITPA treatment

was beneficial to myelination in the IUGR brain, and with unwanted effects of DITPA in controls, it

was essential to also investigate DITPA’s potential therapeutic potential in the cerebellum which is

another important region of the brain with known vulnerability to prenatal insults.

Chapter 4: In this Chapter the impact of the above-mentioned regimen of DITPA treatment in the

cerebellum of IUGR rat pups was explored. This study aimed to determine whether DITPA

Introduction

33

administration (i) impacted cerebellar morphology (ii) promoted OL maturation to restore

myelination, and (iii) did not cause injury or inflammation in the IUGR cerebellum when assessed at

P14. It was hypothesised that DITPA would not impact cerebellar morphology, and would improve

myelination in the cerebellum without causing injury or inflammation. The results of this study

indicate that DITPA administration does not promote myelin recovery (via increased MBP or PLP-

IR), or Olig2-IR OL density in the cerebellum of IUGR pups at P14, and has no negative impact on

cerebellar morphology in IUGR, but may affect Purkinje cell development and cause a slight

inflammatory response in the late developing lobules of the cerebellum. When given to control

animals, the data shows DITPA to reduce the density of BG in the cerebellar ML. These data

collectively support that DITPA administration in IUGR may be more beneficial to the cerebrum than

in the cerebellum. Regardless of the benefit of DITPA, it should not cause unwanted effects on the

individual. Therefore the next step was to determine whether DITPA administration resulted in any

off-target effects on neonatal growth and wellbeing.

Chapter 5: In this Chapter, the effect of DITPA administration on neonatal growth and wellbeing

in IUGR rat pups at P14 was examined. The weight of pups at P1, P7 and P14 was assessed as well

as liver and kidney weights, body morphometry (head and hip circumference, and crown-to-rump

length), body composition measures (including bone density and mineral content, lean tissue mass,

fat mass), thyroid and liver function and cholesterol levels at P14. This study found that DITPA

administration, importantly has no negative off-target effects on neonatal growth, brain or organ

weights, body composition or organ function following IUGR, despite reducing FT4 levels and

showing hepatic thyromimetic activity. However, caution should be taken when administering

DITPA to control pups, as it appears to cause adverse effects when given in a normally functioning

system.

The work presented in this thesis collectively highlights the potential for DITPA to improve

myelination in the IUGR brain, without causing brain injury or adverse off-target physiological

effects, thereby providing valuable preclinical data to inform a clinical trial. However further studies

are required before DITPA can be considered as a therapy in IUGR.

2 General Methodology

34


2.1 Introduction

All of the experiments outlined in this thesis have used a well-established rat model of IUGR. In this

model, first established by Wigglesworth (Wigglesworth, 1964, Wigglesworth, 1974), IUGR is

induced by bilateral uterine vessel ligation (BUVL), which results in placental insufficiency,

mimicking one clinical scenario of IUGR. This model has been used extensively to study brain

development (Lin et al., 1998, Morand et al., 1981, Tashima et al., 2001, Lane et al., 2001a, Baud et

al., 2004, Olivier et al., 2007, Liu et al., 2011, Caprau et al., 2012, Reid et al., 2012, Ke et al., 2014,

McDougall et al., 2017b), injury to organs, as well as long-term metabolic and cardiovascular

outcomes (Gluckman et al., 1996, Jansson and Lambert, 1999, Laker et al., 2012, Ogata et al., 1985,

Peterside et al., 2003, Thompson et al., 2011).

There are a number of reasons this model of IUGR was chosen for our study. Importantly, this model

had been used in our previous short-term DITPA study, in which DITPA was administered from P1

to P6, making it possible to compare and contrast the findings of the present study (i.e. P1 to P13

DITPA administration), with those previously determined by our group. In addition, key time-points

of brain development and in particular myelination and OL development have been well characterised

in the rat, including the Wistar strain (Craig et al., 2003). At the time of BUVL in our model of IUGR

(E18), preOLs have already begun to colonise the brain (Richardson et al., 2006). OL maturation and

myelination occurs mostly after birth in the rat from P1 to 14, and this timeframe is equivalent to the

development of myelin in the human brain at 23 to 40 weeks of GA (Craig et al., 2003, Semple et al.,

2013). In the rat, the development of myelin in the brain at P2 to P5 is similar to the human brain at

23 to 32 weeks GA (Segovia et al., 2008, Wu et al., 2013), and at P7 cerebral myelination is

histologically similar to WM in the human brain at 32 to 34 weeks GA (term = 40 weeks) (Vannucci

and Vannucci, 1997). This model also mimics the clinical effects of IUGR, including reduced body

(Lane et al., 2001b, Olivier et al., 2005, Price et al., 1992, Ogata et al., 1985, Sadiq et al., 1999) and

brain (Sadiq et al., 1999, Ogata et al., 1985, Tashima et al., 2001) weight.

In the brain, TH is essential for OL development, as well as the maturation of the OL lineage from

pre-myelinating OLs to the mature myelinating phenotype (Barres et al., 1994, Rodriguez-Pena,

1999, Billon et al., 2001). Previously our group and others have found that IUGR disturbs the


35

development of a TH transporter protein MCT8, critical for the transport of TH into OLs (Friesema

et al., 2003). In rats, hypothyroidism during brain development can be extremely detrimental to WM

development (Ibarrola and Rodriguez-Pena, 1997, Schoonover et al., 2004, Rodriguez-Pena et al.,

1993). We propose that the synthetic TH analogue, DITPA, which does not require MCT8 to enter

cells, given during a clinically relevant time-frame of WM development can be used to overcome the

myelination deficits caused by the loss of MCT8, and normalise TH transport. The rat IUGR model

allows us to administer DITPA during a relevant window of brain development (P1 to 13 rat = 23 to

40 weeks GA human) (Semple et al., 2013).

2.2 Ethics clearance and animal welfare

Experimental procedures were approved by the Animal Ethics Committee of RMIT University (AEC

project 1702; Appendix 7). All animal handling, use and care follows the National Health and Medical

Research Council of Australia (NHMRC) Code of Practice for the Care and Use of Animals for

Scientific Purposes.

2.3 Animals

48 plug-mated pregnant female Wistar rats (ArcCrl:WI, Wistar outbred) purchased from the Animal

Resource Centre (Perth, Western Australia) at E15 (term = 22) were housed in the RMIT University

Animal Facility (Bundoora) for four days prior to experimentation. This allowed adequate time for

the animals to adapt to the new environment. The pregnant rats were kept in individual cages prior to

surgery, with each rat given unlimited access to food and water. The rats were maintained at a room

temperature of 22C, with a 12-hour light/dark cycle.

2.4 Surgical procedure

2.4.1 Pre-operative preparation

Paracetamol (Children’s Panadol, 2mg/mL in 100mL drinking water; Panadol, Australia) was given

to the rats 24 hours prior to surgery. At E18, rats were randomly allocated to a sham (offspring termed

control) or uteroplacental insufficiency (offspring termed IUGR).

2.4.2 Surgery

Aseptic surgery was conducted at E18 using established techniques. The experimental groups

underwent BUVL surgery (Figure 2.1 A) (Moritz et al., 2009, O'Dowd et al., 2008, Wlodek et al.,

2005, Mazzuca et al., 2010). The control group underwent a sham surgery without the ligation. All


36

instruments, gauze, 0.9% saline and sutures (Vicryl 4-0, Ethicon, USA) were sterilized using an

autoclave. Surgery was performed under aseptic conditions. Anaesthesia was induced using 4%

isoflurane (Isoflo, Abbott, NSW, Australia) in oxygen delivered by nose cone. Sufficient depth of

anaesthesia and the level of unconsciousness were confirmed by the absence of rear foot and eye

reflexes. The dam was then placed in the supine position and administered a maintenance dose of

anaesthetic via a nose cone (2-3% isoflurane in O2) to allow adequate and regular respiration of the

rat. The rat’s lower abdomen was shaved, and thoroughly cleaned with a series of antiseptic solutions

(Chlorohexidene: 0.5% in 70% ethanol, Cenvet, Australia; Betadine: (Cenvet), 70% ethanol). A

sterile drape was placed over the rat, exposing the abdomen, and analgesia was administered

subcutaneously (meloxicam, 5mg/mL, Ilium, USA). Bupivacaine (0.125%, max dose 1-2mg/kg,

Aspen Pharmacare, Australia) was administered at the incision site prior to cutting the skin to provide

local anaesthesia (and pre-emptive analgesia) at the surgical site for 4 to 6 hours. A 2cm midline

abdominal incision was made using sharp end surgical scissors and the cervical end of the uterus

containing foetuses exposed. Sterile gauze soaked in sterile saline was placed on either side of the

incision and the exposed uterus placed carefully on top of this to keep it moist. Both the uterine artery

and veins were ligated using 4-0 braided siliconised silk (Dynek Pty Ltd, Australia) (Figure 2.1

A).The exposed uterus was moistened with sterile saline before being placed back into the body

cavity. Control animals underwent sham surgery using the same technique however without the

uterine vessel ligation. The muscle layer was then sutured using running locked sutures and the

external layer of skin sutured using horizontal mattress sutures (4-0 braided siliconised silk, Dynek

Pty Ltd, Australia). Duration of surgery was 20 to 30 mins. This surgery provides ~30% reduction in

birth weight of offspring (Wigglesworth, 1964).

Figure 2.1 Exposed rat uterus and uterine vessels during bilateral uterine artery ligation (BUVL)

surgery (A). During surgery the uterine horns are exposed, and the cervical end of the uterine vessels are

ligated (sites of ligation marked with yellow crosses), in both the left and right uterine horns. The ligation

restricts ~ 60% of blood flow to the foetuses; the remaining 40% is supplied by the undisturbed ovarian artery.

(B) Representative photograph of an IUGR pup, produced by BUVL surgery (left) compared to a control

surgery pup (right) at P1. Ruler measure is in centimetres (cm). IUGR = intrauterine growth restriction, P1 =

postnatal day 1.


37

2.4.3 End of surgery and post-surgical care

Following surgery, a lubricant (Chloropt, Ceva, Australia) was applied to the eyelids of anaesthetised

rats during recovery, to prevent eye irritation. Rats were returned to the animal room in a new, clean

individual box and placed on a heat-pad overnight to facilitate recovery. Dams fully recovered from

anaesthesia within 15 to 20 min. Once again rats were given access to rat chow and water ad libitum.

Paracetamol (2mg/mL in 100mL drinking water) was given for 2 to 3 days post-surgery. Shredded

paper was provided as nesting material. Rats were monitored closely for the remainder of the day and

then twice daily following surgery. Dams were allowed to deliver naturally; the rat pups were then

housed with their dam and littermates. Rats were checked throughout the day from E21 to 23 for

births.

2.5 Classification of IUGR and control, and size-matching of litters.

Pups were counted and sexed on P1 (to avoid maternal stress immediately after birth). The mean

weight of control pups in our study at P1 (prior to injection and litter matching) was 6.95g (n=100).

Only severe IUGR pups that weighed ≤ 5.61g (6.95g – 2SD [standard deviation]) were used for this

study as described by previous studies (Olivier et al., 2005, Delcour et al., 2012, Tashima et al., 2001).

Control pups from sham surgeries that weighed ≥ 6.27g (6.95g – 1SD) were used for this study as

described by previous studies (Olivier et al., 2007) (Figure 2.1 B). Control litters were on average

larger than IUGR litters, and were therefore size matched to IUGR litter size to ensure an equal

nutritional background across litters. Control and IUGR litters were generated, and underwent

experimental procedures at the same time to control for environmental variables. Only IUGR pups

from BUVL litters were used in these studies, and no IUGR pups from control litters were collected.

2.6 Drug treatment

To determine the effects of longer-term DITPA treatment on the brain, both IUGR and control pups

were injected (i.p) with DITPA (0.5mg/100g, Sigma, USA), or vehicle (equivalent volume of saline)

daily from P1 to P13 (Figure 2.2 A) using a Hamilton syringe (max. volume 50μL) with a disposable

30-gauge needle until P7 and 26-gauge needle from P7 until P13. See Appendix 1 for the protocol

detailing how DITPA is dissolved at this concentration. Thin adhesive film (DuDERM®, ConvaTec,

Australia) was placed onto the site of injection in order to protect the delicate skin of the rat pups

from any irritation. The minimum and maximum weight of rat pups between P1 and P7 was 3 to 12g,

therefore the volume of drug/saline injected was within the range of ~ 10μL to 30μL depending on

pups age/weight. This is well within the maximum allowable volume of 1% of the rat pups body

weight. Dose, timing and route of administration were chosen based on previous rodent studies (Di


38

Cosmo et al., 2009) and human reports (Verge et al., 2012), allowing for cross-species conversion

based on FDA guidelines.

2.6.1 Handling of pups and injection

To minimise possibility of rejection by the mother following handling, recommendations were

followed from the NHMRC Guidelines to Promote Wellbeing of Animals used for Scientific Purposes

Handbook (Part III, Administration of substances, A7). Specifically, all pups were removed from the

litter with their bedding at the same time, gloves were worn to prevent foreign smells being left on

the pups’ skin, and care was taken not to leave any trace of the drugs or blood on the pups’ skin.

Following treatment, the pups were mixed with their original soiled bedding so that they reacquired

the correct smell before being returned to the mother in one go. The injections were made into the

lower right side of the abdomen, alternating the site to avoid bruising or calluses, and the syringe was

drawn back to determine whether any major organs or blood vessels were hit. If this was the case, the

needle was repositioned.

2.6.2 Monitoring after drug treatment

All pups were monitored twice daily (before and after DITPA or saline administration), and included

assessment of body weight (also required to calculate drug dose/volume), changes in normal

behaviour and the appearance of milk spots (to ensure pups were feeding) in the afternoon following

treatment.

2.7 Post-mortem blood and tissue collection

At the completion of the experiment (P14) the dams were euthanised via carbon dioxide inhalation.

Pups were humanely euthanised with an overdose of pentobarbitone sodium (Lethobarb, >100mg/kg

i.p. Virbac, Australia). Blood was collected from each pup, and within litters, both male and female

pups were randomly allocated to one of three brain collection techniques: 1) perfusion fixed and

processed to paraffin wax for histological and immunohistochemical (IHC) analysis (performed in

this thesis), 2) perfusion fixed and frozen into optimal cutting temperature compound (OCT; Tissue

Plus OCT Compound, Scigen, USA) for future IHC analysis (Figure 2.2 E), and 3 un-fixed and snap

frozen in liquid nitrogen and stored at -80°C for future molecular analysis at a later point in time

(Figure 2.2 D). Brains were collected using these different processing techniques to ensure that a

range of analyses could be performed for this thesis and for ongoing studies in our laboratory. Optic

nerves were also carefully collected from male and female animals that were perfused for analysis

using electron microscopy, however were not used in this thesis (Figure 2.2 G).


39

2.7.1 Blood collection

Blood was collected from each pup and placed in Eppendorf tubes with heparin (2µL, 5000 IU in

5mL; Pfizer, Australia) to prevent clotting, then placed on ice. Eppendorf tubes were then centrifuged

at 22° C and 2600rpm for 10 min (Eppendorf 5819 R) to attain plasma for assays (minimum volume

required = 300µL), which was then frozen at -80° C for later analysis using hospital standard plasma

assays (Monash Pathology, Clayton, Victoria, Australia) (Figure 2.2 B) to assess thyroid function

(TSH; plasma FT3 and FT4), plasma liver enzymes alanine transaminase (ALT) and alkaline

phosphatase (ALP) and cholesterol.

2.7.2 Perfusion fixed brain tissue collection

In a fumehood, the euthanised pup was laid on its back and an incision made below the ribcage in the

shape of a ‘v’ ending at the armpits in order to expose the organs. The ribcage was clamped upwards

using haemostats and the diaphragm was cut to reveal the heart. The descending aorta was located

and clamped using a haemostat. A small incision was then made in the right atrium to allow for liquid

to flow out. The ascending circulatory system and brain was then flushed using a 20mL syringe and

a 26-gauge needle filled with 0.9% saline (pH 7.4) inserted into the apex of the heart, until the liquid

exiting the right atrium ran clear. The contents of the syringe was then changed to 4%

paraformaldehyde (PFA, Sharlau, Barcelona, Spain) in 0.1M phosphate buffer (PB; pH 7.4) and the

process repeated in order to fix the brain. Rongeurs were used to gently peel away the skull and

subcutaneous tissue to expose the brain and upper spinal cord. The spinal cord was cleanly cut leaving

at least 1cm still attached to the brain. The brain was gently removed from the skull and post-fixed

for 4 hours in a 50mL container of 4% PFA. The liver and kidneys were weighed then discarded, and

the carcass frozen for later analysis using duel-energy x-ray absorptiometry (DEXA).

2.7.2.1 Perfused brain tissue processed to paraffin wax

Following post-fixation, the olfactory bulbs were removed from the brain using a blade as they were

not relevant to this study. The whole brain (with spinal cord attached) was then weighed, and the

brainstem (pons and medulla only) with the cerebellum attached was dissected from the cerebral

hemispheres, and the hemispheres (also containing the hindbrain) weighed. The cerebellum was

detached from the brainstem at the cerebellar peduncles weighed. The pons, medulla and spinal cord

were dissected and weighed. The cerebellum was divided into left and right hemispheres via a sagittal

cut down the midline (i.e. at the vermis). The left hemisphere of the cerebellum was stored in 4%

PFA (pH 7.4) for Rapid Golgi staining, however this was not part of this thesis. The cerebral

hemispheres, right cerebellar hemisphere, pons, medulla and spinal cord were placed in cassettes and


40

stored in 70% ethanol (maximum 1 day), prior to being processed to paraffin wax at the University

of Melbourne Histology Facility.

2.7.2.2 Perfused brain tissue frozen in OCT

Brain regions were separated and weighed (as outlined in Section 2.7.2.1) and placed into 20%

sucrose in 0.1M PB overnight. The tissue was then placed cutting-face down (ventral side for the

pons, medulla and spinal cord) into peel-away moulds (Grale Scientific Ltd, Australia) which were

carefully filled with OCT, frozen on dry ice and then stored at -80°C for cryo-sectioning and future

analyses by others within our laboratory group.

2.7.3 Fresh snap frozen brain tissue collection

Following euthanasia, the pup’s scalp was opened using a scalpel blade to expose the skull. The brain

was gently removed from the skull as in Section 2.7.2, and then weighed and placed on a petri dish

containing ice to keep it cold. Using the scalpel blade, a sagittal cut was made down the midline of

the brain and spinal cord to separate the right and left hemispheres. The left hemisphere was quickly

placed cutting-face down in a peel-away mould and carefully filled with OCT; the mould was then

wrapped in aluminium foil and frozen on dry ice (until opaque). This tissue was collected for future

studies using laser capture microdissection and RNAseq to assess for differential gene expression.

Using the blade, the remaining hemisphere (right) was separated into the cerebral cortex,

hippocampus, cerebellum, pons, medulla and spinal cord, which were then weighed individually

before being placed into Eppendorf tubes and snap frozen in liquid nitrogen. An incision was made

on the ventral midline of the pup to expose the organs, and the liver, kidneys and adrenals were

removed, weighed and snap frozen in liquid nitrogen. All tissue was stored at -80°C for molecular

analysis see Chapter 5, Section 5.3.5 for assessment of Dio1 mRNA levels in the livers of pups at

P14 and the carcass frozen for later analysis using DEXA scanning (see Section 5.3.3).

2.7.4 Processing of optic nerves

Optic nerves were also collected from pups that were perfusion fixed for the ultrastructural

assessment of myelinated axons. A sharp scalpel was used to make an incision at the anterior of each

eyeball (where the optic nerve attaches), and a fine pair of forceps was then used to grasp the optic

chiasm and gently pull the nerve from the skull. The nerves were post-fixed in 2.5% glutaraldehyde

in 0.1M cacodylate buffer (pH 7.4; sodium cacodylate-trihydrate, Sigma Aldrich Pty Ltd, Australia)

for 4 hours. Following post-fixation, optic nerves were placed in 0.1M cacodylate buffer (pH 7.4) and

transported to the Peter MacCallum Cancer Center Electron Microscopy Facility where they remained


41

in the buffer for a maximum of two days until they were then processed into resin blocks, and semi-

thin sections taken for electron microscopy. Using a JEOL 1010 transmission electron microscope,

myelinated axons within the optic nerve were visualised. Unfortunately the fixation technique used

did not optimally preserve optic nerve tissue, and therefore tissue quality was too poor for parameters

such as myelin sheath thickness, and proportions of myelinated axons to be quantified accurately as

planned. For more information on how optic nerves were processed in preparation for electron

microscopy see Appendix 2.


42

Figure 2.2 Experimental protocol timeline (A) and schematic diagram of tissue collection protocols (B-

G) Pregnant Wistar rats underwent either sham or bilateral uterine vessels (BUVL) surgery at E18 to generate

either IUGR pups, or controls (A). Between P1 and P13 IUGR and control pups received DITPA

(0.5mg/100g/day) or saline treatment (equivalent volume. Rat pups underwent tissue collection at P14. Blood

was collected from all pups and plasma obtained for serum assays (B). Bodies of all pups were frozen for

DEXA scanning and analysis of body and bone composition (C). One male/female pup per litter was

designated to either ‘fresh’ snap frozen tissue collection (D), perfusion to be later processed to paraffin wax,

or frozen in OCT (E). Black stars signify the tissue analysis that was included in this thesis. Optic nerves

(ONs) were collected and processed for electron microscopy (G). BUVL = bilateral uterine vessel ligation,

DEXA = dual-energy x-ray absorptiometry, E = embryonic day, EM = electron microscopy, IF =

immunofluorescence, IHC = immunohistochemistry, LCM = laser capture microdissection, Liquid N2 = liquid

nitrogen, M = molar, OCT = optimal cutting temperature freezing compound, P = postnatal day, P14 =

postnatal day 14, PCR = polymerase chain reaction, PFA = paraformaldehyde, PM = post-mortem.


43

2.8 Tissue histology

2.8.1 Paraffin sectioning of cerebral hemispheres and cerebellum

Please refer to Chapter 3 methods (Section 3.3) for a protocol on sectioning of the cerebral

hemispheres, and Chapter 4 methods (Section 4.3.2) for a protocol on sectioning of the cerebellum.

2.9 Histological staining & analysis

The histological staining protocol below was followed in the staining of both the cerebral hemispheres

and cerebellum.

2.9.1 Haemotoxylin and Eosin (H&E)

Haemotoxylin and Eosin (H&E) staining was performed using a standard procedure. Prior to staining,

sections were dewaxed by clearing through two changes of histolene solvent (Trajan Scientific, Vic,

Australia) and rehydrated through graded solutions of decreasing ethanol concentration to distilled

H2O (dH2O). The tissue was rehydrated and immersed in Mayers Haemotoxylin (Amber Scientific,

WA Australia) for approximately 6 min, to stain the nuclei blue. The sections were then washed under

running tap water, immersed in Scott’s tap water for 1 min, and then submerged in running tap water

for 3 min. The sections were then counterstained with Eosin (1%; Amber Scientific, Australia) for 3

min, which stained the cytoplasm pink. Sections were then submerged in running tap water again for

at least 3 min, until the water ran clear. Sections were then dehydrated through graded solutions of

increasing ethanol concentration and two washes of histolene, before being cover-slipped using DPX

mounting media (DPX, Thermo Scientific Ltd., Vic, Australia).

2.9.2 Analysis of H&E staining

Sections stained with H&E were scanned using Olympus VS120 slide scanner (Olympus, Australia),

and Fiji Software (ImageJ, Version 2.0, National Institute of Health, Maryland, USA) was used to

assess the gross morphology of the cerebellum, with emphasis on the area and width of the cerebellar

layers in cross-section. The presence of haemorrhages, lesions and infarcts were also assessed

qualitatively. Analysis of H&E staining was conducted in the cerebellum only. For an analysis

protocol specific to structures within the cerebellum, please refer to Chapter 4 (Section 4.3.3).

2.10 Immunohistochemical staining

A full outline of final conditions used for each immunohistochemical stain is presented in the

respective Chapters below, in Chapter 3, Table 3.2 for the cerebrum, and in Chapter 4, Table 4.1 for


44

the cerebellum. To avoid procedural variation, and ensure uniform conditions for subsequent analysis,

sections from all four experimental groups were stained simultaneously for each antibody/procedure.

In addition positive controls (P7 rat cerebellum and cerebral cortex) and negative controls (omission

of primary antibody) were performed for each antibody.

A standard immunohistochemical procedure was performed. Following rehydration, sections were

washed in 10% phosphate buffered saline (PBS; 3 x 5 min washes), and antigen retrieval was used to

recover most antigen reactivity in the tissue without increasing background staining. The type of

antigen retrieval was dependent upon the specific protein of interest and included either proteolytic

enzyme-induced epitope retrieval via the use of proteinase K (37˚C for 30mins), or heat induced

epitope retrieval via microwaving in citric acid buffer (pH 6.0, heated to near boiling, simmer 7 mins,

allow to sit at room temperature for 45mins).

Following 3 x 5 min washes in PBS, sections were incubated in 1% hydrogen peroxide (H2O2)

(Merck, Darmstadt, Germany) diluted in PBS for 20 min, in order to block endogenous peroxidase

activity. Following another 3 x 5 min washes in PBS, the sections were incubated with either 4% or

10% bovine serum albumin (BSA, Sigma Aldrich Pty Ltd, NSW, Australia) in 0.1 M PBS (pH 7.4)

to prevent non-specific antibody binding. Sections were then incubated in the appropriate

concentration of primary antibodies diluted in primary diluent (2% BSA in 0.1M PBS + 0.3% Triton

x-100; Sigma Aldrich Pty Ltd, NSW, Australia) overnight at 4˚C.

The following day sections underwent 3 x 5 min PBS washes. The secondary antibodies were diluted

in secondary diluent (2% BSA in 0.1M PBS) and applied to sections for 60min. Sections were washed

in PBS (3 x 5min), and incubated in avidin-biotin peroxidase complex (1:1:200 in PBS; Vectastain

ABC Elite Kit; Vector Laboratories, Burlingame, USA) for 60 min. To enable visualization of the

antibody reactions, 3,3’-diaminobenzadine chromagen (DAB; Mp Biomedicals Australia Pty Ltd,

NSW, Australia; 50mg in 10mL PBS + 3µL 30% H2O2) was added to the slides for approximately 6

to 8 min. Sections were then washed in PBS and then counterstained with Haemotoxylin (Amber

Scientific, WA, Australia), except for slides stained for myelin basic protein (MBP) and glial fibrillary

acidic protein (GFAP). Slides were then dehydrated in graded alcohols, cleared in histolene and

cover-slipped with DPX as described in Section 2.9.1 above.

Immunostained sections were scanned electronically, with a constant light setting using a slide

scanner (Olympus VS120-S5 scanner, Olympus, Vic, Australia); the digital images were visualised

using Fiji (ImageJ, Version 2.0, National Institute of Health, Maryland, USA). For quantitative


45

analyses 3 to 4 sections per region per animal were analysed. Each region of the brain that was stained

immunostained was analysed differently, and was specific to the type of stain. For details on the

analysis of brain regions within the cerebral hemispheres, including the corpus callosum, external

capsule, cortical layer VI, hippocampus and fimbria, please refer to Chapter 3 (Section 3.5). For

details on the immunohistochemical analysis of the cerebellum please refer to Chapter 4 (Section

4.3.5).

Chapter 3

46

3 Impact of DITPA treatment on

myelination and inflammation

in the neonatal IUGR rat

cerebrum

3.1 Introduction

The cerebrum, situated above the cerebellum, is the largest and most anterior region of the brain. It

consists of a left and right hemisphere separated by a midline fissure and contains the cerebral cortex,

the executive functioning centre of the mammalian CNS (Shi et al., 2012). Below the cerebral cortex

are subcortical structures that include the hippocampus, responsible for memory, emotion and spatial

navigation (Wall and Messier, 2001), the fimbria which connects the hippocampus to subcortical

regions, the corpus callosum which relays information between the left and right cerebral

hemispheres, and the external capsule, responsible for connecting different regions of the cortex

(Bloom and Hynd, 2005). The cerebrum is comprised of WM and GM; the overlying cortex is a six-

layered structure, and along with the hippocampus is predominantly comprised of GM. These regions

are rich in neural cell bodies and are essential for processing and integrating information within the

brain. The corpus callosum, external capsule and hippocampal fimbriae are major WM tracts, acting

as ‘information highways’, and are densely packed with myelinated axons. Development of each of

these regions occurs largely prior to birth, and therefore they are vulnerable to prenatal insults such

as the hypoxic/ischaemic injury that often occurs in IUGR, as shown clinically (Takenouchi et al.,

2010, Sizonenko et al., 2006, Kuchna, 1994) and in animal models (Rees et al., 1988, Rees et al.,

1998).

It is known that IUGR babies at an increased risk of neurodevelopmental sequelae (Geva et al., 2006b)

and possess a 10 to 30 fold increased risk of developing cerebral palsy (McIntyre et al., 2013,

MacLennan et al., 2015). A number of clinical neuroimaging studies have found decreased regional

connectivity, which is consistent with decreased WM volume in the cerebrum, including the corpus

callosum and external capsule in IUGR infants at 12 and 18 months of age (Padilla et al., 2014b,

Esteban et al., 2010), changes that persist into childhood and adulthood (Eikenes et al., 2012a).

Chapter 3

47

Decreased volumes of the hippocampus (Lodygensky et al., 2008) and corpus callosum (Egana-

Ugrinovic et al., 2013) have been reported in IUGR infants, as well as compromised development of

the cerebral cortex at post-mortem in IUGR newborns (Dubois et al., 2008). The findings from these

human studies indicate that IUGR negatively impacts cerebral WM and structural development of the

cortex as well as subcortical structures, which may persist into childhood and adulthood.

In IUGR guinea pigs, myelination is reduced in the corticospinal tract at 52 and 62 dg (Nitsos and

Rees, 1990), and in the cerebral cortex at 60 and 62 dg, but not at 1 week of postnatal age (Tolcos et

al., 2011). In the IUGR rat, we have previously shown that myelination is reduced in the corpus

callosum and external capsule at P7 and P14 (Azhan, A., PhD thesis, Monash University, 2019), and

others have reported an overall myelination delay persists into adulthood (Olivier et al., 2005, Reid

et al., 2012). Reduced numbers of pre-OLs have been found in the brains of IUGR rat pups at P7

(Olivier et al., 2005), and reduced numbers of mature OLs in the corpus callosum at P14 (Azhan, A.,

PhD thesis, Monash University, 2019). Given that only mature OLs produce myelin, these results

suggest that the resultant hypomyelination seen in the IUGR brain is a consequence of impaired

maturation of OLs. Based on the findings of these studies, Assessment of myelination will be focused

on in the cerebral layer six (VI) of the cerebral cortex, corpus callosum, external capsule and

hippocampus which are structures vulnerable to the effects of IUGR. Assessment of myelination in

cortical layer VI was chosen, as it was not possible to assess the underlying subcortical WM, due to

the density of staining and because cortical layer VI neurons are the most densely myelinated out of

all the cortical layers (Tomassy et al., 2014). In the hippocampus, the WM in the CA1 region is

examined as this region is important for the recollection of memories (Mueller et al., 2011), and the

CA3 region, essential for encoding new spatial memories (Kesner, 2013). The fimbria, a band of WM

situated along the medial hippocampus that receives large bundles of myelinated fibres from the

hippocampus is also examined.

TH is essential for brain development, and for the maturation of OLs from progenitors to a mature

myelinating phenotype. In the cerebrum, TH is essential for cell migration in the hippocampus and

corpus callosum (Rakic, 1972, Goodman and Gilbert, 2007), as well as the cerebral cortex (Hatten,

1990), where it is required for the correct formation of the six cortical layers (Berbel et al., 2001,

Berbel et al., 1994). A fundamental cellular transporter of TH in the brain is MCT8. However MCT8

protein and mRNA expression is reduced in the IUGR human (Chan et al., 2014) and rat brain (Azhan,

A., PhD thesis, Monash University, 2019) compared to controls. Interestingly, in children with

congenital mutation of MCT8, cerebral myelination is reduced (Lopez-Espindola et al., 2014), and

this suggests that a reduction in MCT8 in the IUGR brain may be compromising cerebral TH

Chapter 3

48

signalling, resulting in reduced myelination. Given that MCT8 is reduced in the IUGR brain (Chan

et al., 2014), treatment with conventional TH may be ineffective, and so alternative TH therapies that

do not rely on MCT8 for cellular uptake need to be tested.

DITPA is a synthetic TH analogue, that readily enter brain cells in MCT8 knockout mice, and

subsequently correct cerebral hypothyroidism (Di Cosmo et al., 2009). A previous study by our group

found that DITPA (0.5mg/100g; i.p.) administered daily from P1 to P6 in IUGR rat pups, a clinical

scenario that mimics an IUGR baby born preterm receiving short-term treatment, promoted myelin

recovery, detectable by P7 in the external capsule (Azhan, A., PhD thesis, Monash University, 2019).

Although promising, this data only provides evidence of the impact of short-term DITPA treatment

on the cerebrum, and although benefits were only seen in the external capsule, perhaps a longer dosing

regimen would further benefit other cerebral and subcortical regions as well.

Therefore the aim of this Chapter was to determine whether DITPA administered to newborn

IUGR rats using a longer-term dosing regimen (daily, from P1 to P13), mimicking a clinical situation

where a preterm IUGR baby is treated with DITPA until term equivalent age (i) restores myelination,

(ii) promotes OL maturation, and (ii) does not cause injury or inflammation, in the cerebral cortex

(layer VI), corpus callosum, external capsule, hippocampus and fimbria. The hypothesis was that

DITPA administration from P1 to 13 will promote and restore OL maturation and myelination within

the cerebral hemispheres of IUGR rat pups when assessed at P14, without causing injury or

inflammation. This study will generate complementary information to our previous short-term

DITPA administration study (Azhan, A., unpublished thesis, 2019) and will inform clinical

translation.

Chapter 3

49

3.2 Methodology

The methodology in this Chapter will commence from the point of paraffin tissue sectioning, and will

focus on the cerebral hemispheres. For information regarding animal welfare, species, surgical

procedure, drug treatment, post-mortem and tissue collection, please refer to Chapter 2 (Sections 2.2

to 2.8).

3.2.1 Overview of animal work

At day 18 of pregnancy (term = 22 days), pregnant Wistar rats underwent BUVL (n= 31 litters) or

sham surgery (n= 16 litters) to generate IUGR or control pups. DITPA (0.5mg/100g; i.p.) or saline

was administered daily from P1 to P13 to IUGR pups (IUGR + DITPA: n= 14 litters, 67 pups; IUGR

+ saline: n= 17 litters, 60 pups) and control pups (control + DITPA: n= 8 litters, 49 pups; control +

saline: n= 8 litters, 47 pups). Not all litters and pups could be used in this study, as the dams ate some

smaller litters, and some BUVL surgeries did not yield IUGR litters or pups. Only males were used

in this study to maintain consistency with our previous short-term DITPA study (Azhan, A., PhD

thesis, Monash University, 2019), and because previous studies examining myelination in IUGR rat

pups did not specify sex (Reid et al., 2012, Olivier et al., 2007). One male pup was randomly selected

from each litter. Therefore the final number of pups per group used in this study is: control + saline,

n= 8; control + DITPA, n= 7; IUGR + saline, n= 8; IUGR + DITPA, n= 8.

3.3 Paraffin sectioning of the cerebral hemispheres

Paraffin embedded P14 rat cerebral hemispheres were sectioned coronally at 8µm using a rotary

microtome (Jung Biocut 2035, Geprufte Sicherhiet, Germany); sections were collected at 3 different

coronal levels - Bregma + 3.00, +1.68 and – 2.68 (Table 3.1) to ensure analysis across the rostro-

caudal extent of the cerebral hemispheres. A total of 45 consecutive sections were cut per animal

from each of the 3 coronal levels (4,185 sections in total; see Figure 3.1 for cutting and staining plan).

This sectioning protocol was applied to capture major WM tracts such as the corpus callosum and

external capsule, cortical WM projections throughout the brain as well as the hippocampus. Sections

were placed in a warm water bath (~40°C) prior to being mounted flat onto slides (Superfrost plus +,

Menzel Glaser, Germany). Three sections per animal were placed onto slides and each section was

separated by 120µm. All tissue-mounted slides were dried overnight at 30 °C prior to histological

and immunohistochemical staining.

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3.4 Immunohistochemical staining of the cerebral hemispheres

Immunohistochemistry was performed using the standard protocol outlined in Chapter 2 (Section

2.12). A number of trials were performed for each antibody to ensure optimal staining quality prior

to conducting the procedure on the actual tissue. For a full outline of final conditions used for each

immunostain performed in the cerebrum, please refer to Table 3.2 To avoid procedural variation, and

ensure uniform conditions for subsequent analysis, sections from all experimental groups were

stained simultaneously for each antibody/procedure. For each animal, 3 sections at each of the 3

coronal levels (9 sections total per animal; see Figure 3.1) were stained with Haematoxylin and Eosin

(for gross morphological assessment e.g. presences of lesions, haemorrhage or regions of pallor) or

immunostained with antibodies directed against MBP, myelin proteolipid protein (PLP),

oligodendrocyte transcription factor 2 (Olig2), adenomatous polyposis coli (APC), glial fibrillary

acidic protein (GFAP) and ionized calcium-binding adaptor molecule 1 (Iba1); sections

immunostained for Olig2, APC and Iba1 were counterstained with Haemotoxylin to visualise the

nuclei of neural cells. In addition, positive controls (P7 rat brain) and negative controls (omission of

primary antibody) were performed for each antibody.

3.5 Immunohistochemical analysis of the cerebral hemispheres

Prior to quantitative assessment, an independent researcher randomly coded sections, so as to allow

blinding to the experimental group. Analyses for each stain were conducted on 9 sections per animal

(3 sections at each of the 3 coronal hemisphere levels, i.e. 3 sections/slide, 120m apart; 31 animals:

31 x 9 sections = 279 sections total analysed for each stain) (see Figure 3.1). The immunostained

slides were digitally scanned using Olympus slide scanning software at 20x magnification, using a

constant light setting (Olympus VS120-S5 scanner, Olympus, Vic, Australia). The digital images

were then visualised and analysed using Fiji software (ImageJ, Version 2.0, National Institute of

Health, Maryland, USA (Schindelin et al., 2012)) and Photoshop software (Photoshop CC, v 19.0,

Adobe Systems Incorporated) to extract images of regions of interest.

Chapter 3

51

Figure 3. 1 Sequence of tissue sectioning and staining. (A) A total of 135 8m-thick sections were cut from

each tissue block, with 45 sections cut at each of the 3 hemisphere levels. (B) Three sections per animal,

separated by 120µm (8µm x 15), per hemisphere level were used for each stain (31 slides per hemisphere level;

93 slides total per stain; 279 slides total including all three hemisphere levels). m = micrometre, APC =

adenomatous polyposis coli, GFAP = glial fibrillary acidic protein, H&E = Haemotoxylin & Eosin, Iba1 =

ionized calcium-binding adaptor molecule 1, MBP = myelin basic protein, Olig2 = oligodendrocyte

transcription factor 2, PLP = myelin proteolipid protein.

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52

3.5.1 Areal coverage (% AC) of MBP-, PLP- and GFAP-immunoreactivity (IR)

MBP and PLP are fundamental proteins within myelin membranes in the CNS compromising around

30% and 50% of the myelin proteins respectively (Baron and Hoekstra, 2010, Werner et al., 2007).

MBP has a role in both myelin formation and stabilisation (Fraser et al., 1989), while PLP plays an

essential role in myelin compaction and wrapping (Klugmann et al., 1997). A reduction in either

MBP- or PLP-IR is an indication of impaired myelination in the brain. GFAP, is a 50kD intermediate

filament protein specific to astrocytes (Eng et al., 1971). Antibodies raised against GFAP are

Table 3.1 Summary of the 3 cerebral hemisphere levels at which analysis was carried out (Bregma: +

3.00, + 1.68, - 2.68), and outline of analysis type carried out at each level and for each stain. Cerebral

hemispheres stained with H&E at each level used for analysis; main structures analysed are outlined or

indicated with text: CA1 = cornu ammonis region 1, CA3 = cornu ammonis region 3,CX = cortex, CC =

Corpus callosum, EC = external capsule, Fim = fimbria, % AC = percentage of areal coverage, APC =

adenomatous polyposis coli, GFAP = glial fibrillary acidic protein, Iba1 = ionized calcium-binding adaptor

molecule 1, MBP = myelin basic protein, Olig2 = oligodendrocyte transcription factor 2.

Chapter 3

53

commonly used to identify astrocytes, with an up-regulation of GFAP indicating an increased density

of astrocytes, due to for example, neuronal injury or hypoxia (Brand and Bignami, 1969).

In layer VI of the cerebral cortex, 3 regions of interest (ROIs; 200µm x 200µm = 40,000 µm2 field

size) were captured from 3 points in the left and right hemispheres using these landmarks: 1st ROI =

above the cingulum; 2nd ROI = above the most lateral extent of the corpus callosum, just prior to the

external capsule, and 3rd ROI = the dorsal part of the external capsule (just before the MBP-IR cortical

fibres end; Figure 3.2).The percentage (%) of a ROI occupied by MBP-IR, PLP-IR and GFAP-IR

(i.e. areal coverage; % AC) was then determined using the ‘threshold analysis tool’ in Fiji software

(threshold analysis). The entire corpus callosum and external capsule (from the left and right

hemisphere) were outlined and defined as ROIs using Photoshop, and the % AC for MBP-IR, PLP-

IR and GFAP-IR were determined using threshold analysis.

In the hippocampus, 4 ROIs were captured in each of the left and right hemispheres. These ROIs

(200µm x 200µm = 40,000 µm2 field size) were positioned laterally against the dorsal edge of the

hippocampus in either the CA1 or CA3 regions, as these regions are both essential for propagation of

information from the hippocampus to the rest of the brain, and more medially, adjacent to the

overlying corpus callosum (one hemisphere: CA1 = 2 ROIs, CA3; 2 ROIs; Both hemispheres = CA1

& CA3 = 4 ROIs each) (see Figure 3.2). Two ROIs were also taken from the fimbria in each

hemisphere (4 ROIs total). For each region (cortical layer VI, corpus callosum, external capsule,

hippocampus and fimbria) the % AC of MBP-IR, PLP-IR and GFAP-IR was averaged per section,

and per animal, and the mean determined for each group.

3.5.2 Projection of MBP- and PLP-IR fibres into the cerebral cortex

The extent to which MBP-IR and PLP-IR fibres projected into the cortex (mm), and the entire cortical

depth of the cortex (mm), were measured at 3 points (1st to 3rd ROI as for % AC) in level matched

sections, using the Fiji ‘line’ and ‘measure’ tool. The percentage of MBP-IR and PLP-IR projection

length to cortex depth at each point was then determined, and this was averaged per section, and per

animal, and the mean determined for each group.

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3.5.3 Areal density of Olig2-, APC- and Iba1-IR cells

Olig2 is a nuclear pan-OL marker in the rodent brain (i.e. marks the entire OL lineage including OL

progenitor cells, pre-OLs, and pre-myelinating and myelinating OLs) (Valerio-Gomes et al., 2018),

while APC is a marker of mature myelinating OLs (Salinas Tejedor et al., 2015). Iba1 is a protein

specific to macrophages and microglia in the CNS and is up-regulated during the activation of these

cells and is therefore an indicator of inflammation or injury to neural cells including OLs (Bennett et

al., 2016).

To examine alterations in the areal density of Olig2-, APC- and Iba1-IR cells between groups,

analysis was carried out in cortical layer VI, hippocampus and fimbria using the same ROIs (200µm

x 200µm = 40,000 µm2 field size) as described for MBP-IR and PLP-IR analysis in Section 3.5.1

above. However, instead of using a threshold analysis and % AC, positively stained Olig2-, APC-

and Iba1-IR cells were counted within each ROI and the number divided by the ROI field size to

determine areal density (cells/mm2). In the corpus callosum, counts were conducted in 4 ROIs

(200µm x 200µm = 40,000 µm2 field size) spaced randomly and evenly along the corpus callosum in

each section. For each region (cortical layer VI, hippocampus, fimbria and corpus callosum) data was

averaged per section, the 3 sections were averaged to give cells/mm2 per animal, and means were

determined for each group. The percentage of mature APC-IR OLs within the entire OL population

(Olig2-IR) was calculated (APC-IR (cells/mm2)/ Olig2-IR (cells/mm2)) and averaged per animal.

Figure 3.2 Coronal section of P14 rat cerebrum stained with MBP Quantitative analyses of

immunohistochemical staining were performed with 6 fields in the cerebral cortex (layer VI; black boxes), the

entire corpus callosum (labelled CC; central), the external capsule (yellow trace),4 fields in the hippocampal

CA1 region (green boxes), and 4 fields in the CA3 region (pink boxes) and 4 in the fimbria (blue boxes) per

coronal level (level 3 pictured here). Square regions of interest (to size) = 200µm x 200µm = 40,000 µm2 field

size. CC = Corpus callosum, EC = external capsule.

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3.6 Statistical analysis

All statistical analyses were performed using the Graphpad Prism statistical software (Graphpad

Software, Version 8.0, CA, USA). Prior to analyses, outliers were removed using the Grubb’s test to

determine a significant outlier (p < 0.05). Data were checked for normal distribution, and if found to

not be normally distributed, the appropriate non-parametric test was used instead (Mann-Whitney U

test, instead of unpaired t-test; Friedman test instead of two-way ANOVA). A two-way ANOVA was

used, with a post-hoc pairwise comparison to identify between group differences, and a Bonferroni

correction was used for multiple post-hoc comparisons between experimental groups. Data were

considered significant if p < 0.05. All main effects data are presented in text as ratio of residual

variances (F) and statistical significance (p-value). All post-hoc data are presented as [mean

comparative difference (95% confidence interval); and p-value], and graphs are represented as Mean

standard error of the mean (SEM). All two-way ANOVA results are tabulated in Appendix 3 as

Mean SEM.

Chapter 3

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Table 3. 2 Immunohistochemistry - optimised antigen retrieval, blocking protocol and antibody concentrations in the cerebrum.

Abbreviations: Ab, antibody; APC, adenomatous polyposis coli; BSA, bovine serum albumin; cat, catalogue number; DAB, diaminobenzidine; GFAP, glial fibrillary

acidic Protein; H2O2, hydrogen peroxide; Iba1, Ionized calcium-binding adaptor molecule 1; M, molar; MBP, Myelin basic protein; Olig2, oligodendrocyte

transcription factor-2; PBS, pH; Power of the hydrogen atom; phosphate buffered saline; PLP, myelin proteolipid protein.

Protein Staining for Antigen Retrieval & block 1°Ab (source); conc. 2°Ab (source); conc. Complex

binding to

secondary

and DAB

[Complex][DA

B]

MBP Myelin in the CNS Heat in 0.01M citrate buffer (pH

6.0) for 9 min, then decreasing

power to maintain simmer for 7

min; 4% BSA in PBS

Rat anti-MBP,

(Millipore cat:

MAB386); 1:250

Biotinylated rabbit anti-rat,

(Millipore); 1:200

Avidin-biotin

complex

DAB (in stable

peroxide

buffer)

50μl A + 50μl B

in 10 mL PBS;

1 DAB tablet in

10mL dH2O +

3μl H2O2

PLP Myelin in the CNS Heat in 0.01M citrate buffer (pH



min; 4% BSA in PBS

Mouse anti- PLP

(Millipore cat:

MAB388); 1:500

Biotinylated goat anti -

mouse, (Vector cat:

VEBA9200); 1:200

Olig2 Entire OL lineage Heat in 0.01M citrate buffer (pH



min;

4% BSA in PBS

Rabbit anti-Olig2,

(Millipore cat:

MABN50); 1:500

Biotinylated goat anti-

rabbit,

(Vector cat: VEBA1000);

1:200

APC Mature OLs Heat in 0.01M citrate buffer (pH



min; 10% BSA in PBS

Mouse anti- APC – CC1

(Millipore); 1:500


mouse, (Vector cat:

VEBA9200); 1:200

GFAP Bergmann glial

cells & astrocytes

1:500 Proteinase K in 37°C for 30

min; 4% BSA in PBS

Rabbit anti-GFAP,

(DAKO cat: 0334):

1:500


rabbit,


1:200

Iba1 Microglia &

macrophages in

CNS

Heat in 0.01M citrate buffer (pH



min;

4% BSA in PBS

Rabbit anti-Iba-1,

(Wako cat: 019-19741);

1: 1000


rabbit,


1:200

Chapter 3

57

3.7 Results

3.7.1 Myelination and oligodendrocytes

3.7.1.1 Areal coverage (% AC) of MBP-IR in cortical layer VI

Two-way ANOVA analysis showed no interaction between group and treatment on the percentage of

area covered (% AC) by MBP-IR in cortical layer VI, but there was a main effect of group and

treatment on % AC (group: F1/256 = 17.74; p < 0.0001; treatment: F1/256 62.95; p < 0.0001). Post-hoc

analysis showed that % AC of MBP-IR in cortical layer VI was not different in IUGR + saline

compared to control + saline pups (Figure 3.3 A). There was an increase in % AC of MBP-IR in

IUGR + DITPA compared to IUGR + saline pups [8.39 (3.70, 13.04); p < 0.0001; Figure 3.3 A], and

in control + DITPA compared to control + saline pups [11.48 (6.70, 16.26); p < 0.0001; Figure 3.3

A]. There was a reduction in % AC of MBP-IR in IUGR + DITPA compared to control + DITPA

pups [-6.82 (-11.49, - 2.14); p = 0.0008; Figure 3.3 A].

3.7.1.2 Percentage of cerebral cortex occupied by MBP-IR fibres

Two-way ANOVA analysis showed no interaction between group and treatment on the percentage of

cerebral cortex occupied by MBP-IR fibres, however there was a main effect of group (F1/68 = 8.68,

p = 0.004) and treatment (F1/68 = 10.57, p = 0.002). Post-hoc analysis showed a decrease in the

percentage of the cerebral cortex occupied by MBP-IR fibres in IUGR + DITPA compared to IUGR

+ saline pups [6.23 (-11.84, -0.62); p = 0.02; Figure 3.3 B], and no difference between any other

groups.

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3.7.1.3 Areal coverage (% AC) of MBP-IR in the corpus callosum and external

capsule

Two-way ANOVA analysis showed no interaction between group and treatment on % AC of MBP-

IR in the corpus callosum (Figure 3.4 A) and external capsule (Figure 3.4 B), and no main effects of

group or treatment.

Figure 3.3 Areal coverage (% AC) of MBP-IR (A) in cortical layer VI, and proportion (%) of cerebral

cortex depth containing MBP-IR fibre projections (B) at P14 in control and IUGR pups treated with

DITPA or saline. Representative photomicrographs of MBP-IR in the cerebral cortex (layer VI) of a control

+ saline (C), control + DITPA (D), IUGR + saline (E) and IUGR + DITPA (F) pup show increased % AC by

MBP-IR in IUGR + DITPA pups compared to IUGR + saline, and control + DITPA pups compared to control

+ saline. * p <0.05, *** p < 0.001, **** p < 0.0001. Data analysed using a two-way ANOVA with Bonferroni

correction. Values presented as Mean ± SEM. Pup numbers: control + saline: n= 8; control + DITPA: n= 7;

IUGR + saline: n= 8; IUGR + DITPA: n= 8. All animals are male. Scale bar = 0.05 mm.

Chapter 3

59

3.7.1.4 Areal coverage (% AC) of MBP-IR in the hippocampus and fimbria

Two-way ANOVA analysis showed no interaction between group and treatment on the % AC of

MBP-IR in the CA1 or CA3 regions of the hippocampus, or in the fimbria. There was a main effect

of group in the hippocampal CA1 (F1/58 = 55.58, p < 0.0001) and CA3 regions (F1/57 = 19.74, p <

0.0001), and an effect of treatment in the fimbria (F1/25 = 163.1, p < 0.0001). Post-hoc analysis

showed reduced % AC of MBP-IR in the CA1 and CA3 regions of the hippocampus in IUGR + saline

compared to control + saline pups [CA1: 3.78 (1.26, 6.31); p = 0.0012; CA3: 3.61 (0.21,7.01); p =

0.034; Figure 3.4 C, D], but no difference in the fimbria (Figure 3.4 E). There was an increase in %

AC of MBP-IR in the fimbria of control + DITPA compared to control + saline pups [-33.82 (-44.81,-

22.83) p < 0.0001], and IUGR + DITPA compared to IUGR + saline pups [-34.04 (-44.58, -23.51); p

< 0.0001; Figure 3.4 E]. In IUGR + DITPA compared to control + DITPA pups, % AC of MBP-IR

was reduced in the hippocampal CA1 and CA3 regions [CA1: 6.47 (3.85, 9.1); p < 0.0001; CA3: 4.67

(1.10, 8.25); p = 0.006; Figure 3.4 C, D].

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Figure 3.4 Areal coverage (% AC) of MBP-IR in the corpus callosum (A), external capsule (B),

hippocampal CA1 region(C), hippocampal CA3 regions (D), and fimbria (E) at P14 in control and

IUGR pups treated with DITPA or saline. Representative photomicrographs of MBP-IR in the fimbria of

a control + saline (F), control + DITPA (G), IUGR + saline (H) and IUGR + DITPA (I) pup show increased

% AC of MBP-IR in IUGR + DITPA pups compared to IUGR + saline, and control + DITPA pups compared

to control + saline. * p < 0.05, ** p < 0.01, **** p < 0.0001. Data analysed using a two-way ANOVA with

Bonferroni correction. Values presented as Mean ± SEM. Pup numbers: control + saline: n= 8; control +

DITPA: n= 7; IUGR + saline: n= 8; IUGR + DITPA: n= 8. All animals are male. Scale bar = 0.05 mm.

Chapter 3

61

3.7.1.5 Areal coverage of PLP-IR in cortical layer VI

Two-way ANOVA results showed no interaction between group and treatment on the % AC of PLP-

IR in cortical layer VI, however there was a main effect of group (F1/219 = 73.83, p < 0.0001), and a

main effect of treatment (F1/219 = 5.49, p = 0.02). Post-hoc analysis showed reduced % AC of PLP-

IR in IUGR + saline compared to control + saline pups [-13.05 (-17.81, -8.30); p < 0.0001], and in

IUGR + DITPA pups compared to control + DITPA pups [-9.22 (-14.22, -4.22); p < 0.0001; Figure

3.5 A].

3.7.1.6 Percentage of cerebral cortex occupied by PLP-IR fibres

Two-way ANOVA results showed no interaction between group and treatment of the percentage of

cerebral cortex occupied by PLP-IR fibres, and no main effect of group or treatment (Figure 3.5 B).

3.7.1.7 Areal coverage (% AC) of PLP-IR in the corpus callosum and external

capsule

Two-way ANOVA results showed no interaction between group and treatment on the % AC of PLP-

IR in the corpus callosum and external capsule. There was a main effect of group and treatment on

the % AC of PLP-IR in the corpus callosum (group: F1/51 = 12.09, p = 0.48; treatment: F1/51 = 34.00,

p < 0.0001), and the external capsule (group: F1/118 = 81.09, p < 0.0001; treatment: F1/118 = 10.19, p

= 0.002). Post-hoc analysis showed reduced % AC of PLP-IR in the external capsule of IUGR +

saline compared to control + saline pups [-13.12 (-18.38, - 7.87); p < 0.0001; Figure 3.6 B]. The %

AC of PLP-IR was reduced in the corpus callosum [-12.66 (-25.18, - 0.16); p = 0.04; Figure 3.6 A]

Figure 3.5 Percentage of area covered by PLP-IR (A) in the cortex (layer VI), and cortical projection

length (B) at P14 in control and IUGR pups treated with DITPA or saline. Data analysed using a two-way

ANOVA with Bonferroni correction. Values presented as Mean ± SEM. **** p < 0.0001. Pup numbers: control

+ saline: n= 8; control + DITPA: n= 7; IUGR + saline: n= 8; IUGR + DITPA: n= 8. All animals are male.

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and external capsule [-7.62 (-12.88, - 2.36); p = 0.001; Figure 3.6 B] of IUGR + DITPA when

compared to IUGR + saline pups, and in control + DITPA compared to control + saline pups in the

external capsule [-8.69 (-14.04, -3.33); p = 0.0002; Figure 3.6 B]. In IUGR + DITPA compared to

control + DITPA pups the % AC of PLP-IR was reduced in the corpus callosum [-13.59 (-25.85, -

1.32); p = 0.02; Figure 3.6 A] and in the external capsule [-12.06 (-17.41, -6.71); p < 0.0001; Figure

3.6, B].

3.7.1.8 Areal coverage (% AC) of PLP-IR in the hippocampus and fimbria

Two-way ANOVA results showed an interaction between group and treatment on the % AC of PLP-

IR in the CA1 region of the hippocampus (F1/23 = 5.65, p = 0.026). Main effects of group and

treatment were also shown in the CA1 region (group: F1/23 = 30.76, p < 0.0001; treatment: F1/23 =

7.02, p = 0.014), and an effect of group only in the CA3 region (F1/22 = 12.26, p = 0.0020) and

fimbria (F1/23 = 30.02, p < 0.0001). Post-hoc analysis found that in IUGR + saline compared to

control + saline pups the % AC of PLP-IR was reduced in the CA1 region of the hippocampus [9.70

(4.57, 14.83) p < 0.0001] and in the fimbria [11.30 (4.44, 18. 15); p = 0.0005; Figure 3.6 C, E] . There

was a reduction in % AC of PLP-IR in the CA1 region of control + DITPA compared to control +

saline pups [6.15 (0.71, 11. 60) p = 0.021; Figure 3.6 E], and in the fimbria of IUGR + DITPA

compared to control + DITPA pups [6.63, (0.14, 13.12); p = 0.04; Figure 3.6, E]. There was no

difference in the % AC of PLP-IR in the hippocampal CA3 region between groups (Figure 3.6 D).

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Figure 3.6 Areal coverage (% AC) of PLP-IR in the corpus callosum (A, B), external capsule (C, D),

hippocampal CA1 (E) and CA3 (F) regions and fimbria (G) at P14 in control and IUGR pups treated

with DITPA or saline. Representative photomicrographs of PLP-IR in the external capsule of a control + saline (F), control + DITPA (G), IUGR + saline (H) and IUGR + DITPA (I) pup show decreased % of PLP-IR AC in IUGR + DITPA pups compared to IUGR + saline, and control + DITPA pups compared to control + saline Data

analysed using a two-way ANOVA with Bonferroni correction. Values presented as Mean ± SEM. * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001. Pup numbers: control + saline: n= 8; control + DITPA: n= 7; IUGR + saline: n= 8; IUGR + DITPA: n= 8. All animals are male.

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3.7.1.9 Areal density of Olig2-IR OLs in cortical layer VI

Two-way ANOVA results showed an interaction between group and treatment on the areal density

of Olig2-IR OLs in cortical layer VI (F1/274 = 11.14, p = 0.001); there was no main effect of group or

treatment. Post-hoc analysis revealed that areal density of Olig2-IR OLs was decreased in cortical

layer VI in IUGR + saline pups compared to control + saline pups [-110 (-189, - 31); p = 0.0016],

and in control + DITPA pups compared to control + saline [-96.94 (-178.8, -15.09); p = 0.01] (Figure

3.7, A). There was also no difference in density of Olig2-IR OLs in IUGR + DITPA compared to

IUGR + saline pups (Figure 3.7 A).

3.7.1.10 Areal density of Olig2-IR oligodendrocytes in the corpus callosum

Two-way ANOVA results showed no interaction between group and treatment on the areal density

of Olig2-IR OLs in the corpus callosum, however there was a main effect of treatment (F1/58 = 10.87,

p = 0.002). Post-hoc analysis showed that the areal density of Olig2-IR OLs was increased in the

corpus callosum of IUGR + DITPA compared to IUGR + saline pups [534 (85.86, 982); p = 0.01;

Figure 3.7, B]; there was no difference between IUGR + saline and control + saline pups, or between

control + DITPA and control + saline pups (Figure 3.7 B).

3.7.1.11 Areal density of Olig2-IR oligodendrocytes in the hippocampus and fimbria


of Olig2-IR oligodendrocytes in the fimbria (F1/27 = 4.49, p = 0.04). Post-hoc analysis showed that

there was no difference in areal density of Olig2-IR OLs in the CA1 region (Figure 3.7 C), CA3

region (Figure 3.7 D) or the fimbria (Figure 3.7 E) between groups.

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3.7.1.12 Areal density of APC-IR OLs in cortical layer VI

Two-way ANOVA analysis showed an overall interaction between group and treatment on the areal

density of APC-IR OLs in layer VI of the cerebral cortex (F1/100 = 9.4, p = 0.003) and a main effect

of group on areal density of APC-IR OLs (F1/100 = 7.10, p = 0.009). Post-hoc analysis revealed a

reduction the areal density of Olig2-IR cells in IUGR + saline compared to control + saline pups [-

196 (-327.5, -65.76); p = 0.0006]. The density of Olig2-IR cells was also reduced in control + DITPA

Figure 3.7 Areal density of Olig2-IR oligodendrocytes in the cortical layer VI (A), corpus callosum (B),

hippocampal CA1 region (C), hippocampal CA3 region (D), and fimbria (E) at P14 in control and IUGR

pups treated with DITPA or saline. Representative photomicrographs of Olig2-IR in the corpus callosum of

a control + saline (F), control + DITPA (G), IUGR + saline (H) and IUGR + DITPA (I) pup show increased

Olig2-IR OL density in IUGR + DITPA pups compared to IUGR + saline. Data analysed using a two-way

ANOVA with Bonferroni correction. Values presented as Mean ± SEM. * p < 0.05, ** p < 0.01. Pup numbers:

control + saline: n= 8; control + DITPA: n= 7; IUGR + saline: n= 8; IUGR + DITPA: n= 8. All animals are

male. Scale bar = 0.05 mm.

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compared to control + saline pups [-149 (-278, -19.7); p = 0.02; Figure 3.8 A]. There was no

difference in the density of Olig2-IR OLs between IUGR + DITPA and IUGR + saline pups (Figure

3.8 A).

3.7.1.13 Areal density of APC-IR OLs in the corpus callosum

Two-way ANOVA analysis showed an interaction between group and treatment in the corpus

callosum (F1/53 = 5.34, p = 0.02), and a main effect of group on the areal density of APC-IR OLs (F1/53

= 28.75, p < 0.0001). Post-hoc analysis showed a decrease in the areal density of APC-IR OLs in the

corpus callosum of IUGR + saline compared to control + saline pups [-422.6 (-630.6 – 214.7); p <

0.0001], but there was no difference between IUGR + saline and IUGR + DITPA pups (Figure 3.8

B).

3.7.1.14 Areal density of APC-IR OLs in the hippocampus and fimbria


of APC-IR OLs in the CA1 region of the hippocampus and the fimbria (CA1: F1/25 = 5.04, p = 0.03;

Fimbria: F1/25 = 4.91, p = 0.04). There was a main effect of group on the areal density of APC–IR

cells in the CA1 and CA3 regions (CA1: F1/25 = 6.40, p = 0.02; CA3: F1/25 = 5.73, p = 0.02). Post-hoc

analysis showed a significant reduction in the areal density of APC-IR OLs in the CA1 [375.5 (74.3,

676.7); p = 0.009; Figure 3.8 C] and CA3 region [378.1 (38.19, 718.1); p = 0.02; Figure 3.8 D] of the

hippocampus, and in the fimbria [858.3 (13.09, 17.04); p = 0.045; Figure 3.8 E] of IUGR + saline

compared to control + saline pups; there was no difference in IUGR + saline compared to IUGR +

DITPA pups in any regions (Figure 3.8 A- E).

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3.7.1.15 Percentage (%) of mature APC-IR OLs in cortical layer VI

Two-way ANOVA analysis showed an interaction between group and treatment on the percentage of

mature APC-IR OLs within the overall Olig2-IR OL population in cortical layer VI (F1/284 = 31.74, p

< 0.0001). There was a main effect of group and treatment on the percentage of mature APC-IR OLs

Figure 3.8 Areal density of APC-IR oligodendrocytes in the cortical layer VI (A), corpus callosum (B),

hippocampal CA1 region (C), hippocampal CA3 region (D), and fimbria (E) at P14 in control and

IUGR pups treated with DITPA or saline. Representative photomicrographs of APC-IR in cortical layer

VI of a control + saline (F), control + DITPA (G), IUGR + saline (H) and IUGR + DITPA (I) pup show

decreased APC-IR OL density in IUGR + saline pups compared to control + saline (F vs. H) and control +

DITPA compared to control + saline (F vs. G). Data analysed using a two-way ANOVA with Bonferroni

correction. Values presented as Mean ± SEM. * p < 0.05, ** p < 0.01, *** and ****p < 0.0001. Pup numbers:

control + saline: n= 8; control + DITPA: n= 7; IUGR + saline: n= 8; IUGR + DITPA: n= 8. All animals are

male.

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(group: F1/284 = 6.27, p = 0.01; treatment: F1/248 = 11.78, p = 0.0007). Post-hoc analysis showed a

decrease in the percentage of mature APC-IR OLs in IUGR + saline compared to control + saline

pups [-27.46 (-40.13, - 14.78); p < 0.0001]. There was a lower percentage of mature OLs in control

+ DITPA pups compared to control + saline pups [-30.58 (-43.26, -17.91); p < 0.0001; Figure 3.9 A];

there was no difference in IUGR + DITPA compared to IUGR + saline pups (Figure 3.9 A).

3.7.1.16 Percentage (%) of mature APC-IR OLs in the corpus callosum

Two-way ANOVA results showed an interaction between group and treatment on the percentage of

mature APC-IR OLs within the overall Olig2-IR OL population in the corpus callosum (: F1/60 = 6.87,

p = 0.01). There were no main effects of group or treatment. Post-hoc analysis showed a decreased

percentage of mature APC-IR OLs in IUGR + saline pups compared to control + saline [-21.74 (-

43.20, -0.38); p = 0.04; Figure 3.9 B). There was no difference in the percentage of mature APC-IR

OLs in the corpus callosum between IUGR + DITPA and IUGR + saline groups (Figure 3.9 B).

3.7.1.17 Percentage (%) of mature APC-IR OLs in the hippocampus and fimbria

Two-way ANOVA analysis found no interaction between group and treatment on the percentage of

mature APC-IR OLs within the overall Olig2-IR OL population in regions CA1 and CA3 of the

hippocampus, or in the fimbria; there was a main effect of group in the hippocampal CA1 and CA3

regions (CA1: F1/28 = 7.78, p = 0.009; CA3: F1/28 = 6.14, p = 0.02). Post-hoc analysis revealed a

decrease in the percentage of mature APC-IR OLs in the fimbria of control + DITPA pups compared

to control + saline pups [-29.64 (-56.90, -2.38); p = 0.03; Figure 3.9 E], but no difference in IUGR +

DITPA pups compared to IUGR + saline, or in IUGR + saline compared to control + saline pups;

there were no difference between groups in the hippocampal CA1 and CA3 regions (Figure 3.9 C,

D).

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3.7.2 Inflammation

3.7.2.1 Areal density of Iba1-IR microglial

Two-way ANOVA analysis showed no interaction between group and treatment on the areal density

of Iba1-IR microglia (cells/mm2) in cortical layer VI, the corpus callosum, hippocampal CA1 and

CA3 regions or in the fimbria, and no main effects of group or treatment (Figure 3.10 A-E).

Figure 3.9 Proportion of mature APC-IR OLs to total OL lineage (APC:Olig2) in the cortical layer VI

(A), corpus callosum (B), hippocampal CA1 region (C), hippocampal CA3 region (D), and fimbria (E) at

P14 in control and IUGR pups treated with DITPA or saline. Data analysed using a two-way ANOVA with

Bonferroni correction. Values presented as Mean ± SEM. * p < 0.05, **** p < 0.0001. Pup numbers: control + saline: n= 8; control + DITPA: n= 7; IUGR + saline: n= 8; IUGR + DITPA: n= 8. All animals are male.

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Figure 3.10 Density of Iba1-IR microglia in the cortex (layer VI; A), corpus callosum (B), CA1 (C) and

CA3 (D) regions of the hippocampus and fimbria (E) at P14 in control and IUGR pups treated with

DITPA or saline. Representative photomicrographs of GFAP-IR in cortical layer VI of a control + saline (F),

control + DITPA (G), IUGR + saline (H) and IUGR + DITPA (I) pup show no difference in GFAP-IR astrocyte density between groups. Data analysed using a two-way ANOVA. Values presented as Mean ± SEM. Pup numbers: control + saline: n= 8; control + DITPA: n= 7; IUGR + saline: n= 8; IUGR + DITPA: n= 8. All animals are male.

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3.7.2.2 Areal coverage (% AC) of GFAP-IR

Two-way ANOVA analysis showed no interaction between group and treatment on the % AC of

GFAP-IR astrocytes in cortical layer VI, corpus callosum, external capsule, hippocampal CA1 and

CA3 regions or fimbria. There were no main effects of group or treatment in any of these regions

(Figure 3.11).

0.05mm

Figure 3.11 Percentage of area covered by GFAP-IR astrocytes in the cortex (layer VI; A), corpus

callosum (B), external capsule (C), CA1 (D) and CA3 (E) regions of the hippocampus and fimbria (F) at

P14 in control and IUGR pups treated with DITPA or saline. Representative photomicrographs of GFAP-

IR in cortical layer VI of a control + saline (G), control + DITPA (H), IUGR + saline (I) and IUGR + DITPA

(J) pup show no difference in GFAP-IR astrocyte density between groups. Data analysed using a two-way

ANOVA. Values presented as Mean ± SEM. Pup numbers: control + saline: n= 8; control + DITPA: n= 7;

IUGR + saline: n= 8; IUGR + DITPA: n= 8. All animals are male.

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3.8 Discussion

3.8.1 Overview

This is the first study to examine the structure of the neonatal cerebral hemispheres in response to

daily, longer-term DITPA administration (P1 to 13) in an IUGR rat model. As described previously,

analyses were carried out at three hemispheric levels to account for the developmental profile of the

cerebral WM and then averaged per animal. The major findings of this study are:

In IUGR + saline compared to control + saline pups there was significantly reduced (i) % AC of

MBP-IR in the hippocampus, (ii) % AC of PLP–IR in cortical layer VI, external capsule,

hippocampus and fimbria, (iii) areal density of all OLs (Olig2-IR; cells/mm2) in cortical layer VI, (iv)

areal density of APC-IR mature OLs, in cortical layer VI, corpus callosum, hippocampus and fimbria,

and (iv) percentage of APC-IR mature OLs within the entire population of OLs in cortical layer VI

and in the corpus callosum. There was (v) no difference in the areal density of microglia (Iba1) and

the % AC of astrocytes (GFAP) in any of the regions examined.

In IUGR + DITPA compared to IUGR + saline pups there was significantly (i) increased % AC of

MBP-IR in cortical layer VI and the fimbria, (ii) increased density of Olig2-IR OLs in the corpus

callosum, (iii) reduced percentage of cerebral cortex occupied by MBP-IR projections, and (iv)

reduced % AC of PLP–IR in the corpus callosum and external capsule.

In control + DITPA compared to control + saline pups there was significantly (i) increased % AC of

MBP-IR in cortical layer VI and the fimbria, (ii) reduced % AC of PLP-IR in the external capsule

and the CA1 region of the hippocampus, (iii) reduced areal density of Olig2-IR OLs and (iv) reduced

areal density of APC-IR OLs in cortical layer VI, as well as (v) reduced percentage of APC-IR mature

OLs within the entire population of OLs in cortical layer VI and fimbria.

3.8.2 Effects of daily DITPA administration on white matter development

3.8.2.1 Myelin proteins

MBP and PLP are two of the main myelin proteins in the CNS, and are integral to the structure of the

myelin sheath, with MBP and PLP making up roughly 30% and 50% of myelin proteins respectively

(Baron and Hoekstra, 2010, Werner et al., 2007). The immunohistochemical staining of these two

proteins was used to determine the overall effects of DITPA on the cerebral WM in newborn IUGR

rat pups.

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MBP-IR

This study found that % AC of MBP-IR in the corpus callosum, external capsule, and cortical layer

VI was not affected by IUGR, but there was a reduction in the hippocampus (CA1 and CA3 regions).

This finding was unexpected, as a previous study by our group using an IUGR rat model,

demonstrated that % AC of MBP-IR was significantly reduced in the corpus callosum and external

capsule at P7 and P14, and that DITPA administration (0.5mg/100kg/day i.p. from P1 to P6) corrected

the hypomyelination by P7 in the external capsule (Azhan, A., PhD thesis, Monash University, 2019).

A reduction in myelin proteins in the brain of IUGR infants has been reported at post-mortem (Chase

et al., 1972), as well as in other animal models (Reid et al., 2012, Tolcos et al., 2011, Azhan, 2019),

and therefore a more pronounced effect of IUGR on cerebral myelination was expected in this study.

The hippocampal CA1 region is essential for encoding long-term memory and spatial recognition

(Duvernoy, 2005), and the CA3 is fundamental for encoding short-term memory and spatial

information (Cherubini and Miles, 2015), therefore disruptions to MBP-IR in these regions may have

an impact on memory processing capacity in the hippocampus. The hippocampus has known

vulnerability to the effects of hypoxia and under-nutrition, both known to occur in IUGR

(Lodygensky et al., 2008, Sizonenko et al., 2006), and disruptions to hippocampal development in

IUGR is seen in guinea pigs (Mallard et al., 1999, Mallard et al., 2000). This vulnerability could

explain why a decrease in % AC of MBP-IR was seen in the hippocampus of IUGR pups in this study.

It is still not clear why a reduction in % AC of MBP-IR was not seen in several brain regions of the

IUGR pups in this study at P14, One possibility could be that the stress of daily handling and injecting

of pups (with either saline or DITPA) for an extended period of time (from P1 to 13, rather than P1

to P6 as in (Azhan, A., PhD thesis, Monash University, 2019) is having an effect on myelination in

the control pups as well. It is known that cortisol released during a stress response delays myelination

of major WM tracts in the brain and spinal cord (Bohn and Friedrich, 1982, Antonow-Schlorke et al.,

2009, Shields et al., 2012). Our pups underwent roughly 2 to 3 min of physical handling per day (e.g.

weighing, marking), as well as the physical and psychological stress of daily intraperitoneal injections

and being briefly separated from their mother during this process. Stress is likely having an effect on

both control and IUGR pups, and therefore a level of reduced myelination would theoretically be

expected in both. However, could it be possible that OLs in IUGR pups are more resilient to the

effects of cortisol than controls as they have developed under the stress of IUGR conditions? If this

were the case, myelination would more greatly affected in control pups compared to IUGR pups in

response to stress, thus lowering their MBP-IR levels to that seen in IUGR pups. Indeed, late

progenitor OLs are more likely to be killed by hypoxic-ischemic insult, like that occurring in IUGR,

and that the surviving progenitors and mature OLs are more resilient after the insult, thus

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compensating for the loss in numbers by increasing their density and accelerating maturation (Back

et al., 2001, Back et al., 2002). In our model, could resilience in the surviving OLs in the cerebrum

of IUGR pups, compared to controls, underlie the lack of difference in MBP-IR between IUGR and

control pups? Future studies should aim to study the density and re-myelinating potential of OLs in

the IUGR rat brain, with and without exposure to daily handling and compare this to OLs in the non-

IUGR rat brain in order to answer this question.

In the present study, following DITPA administration % AC of MBP-IR was increased in cortical

layer VI of both IUGR and control pups compared to saline, and there was a concurrent decrease in

the percentage of cerebral cortex occupied by MBP-IR projections in IUGR pups given DITPA

compared to those given saline. This suggests that DITPA treatment is resulting in denser MBP-IR

along fibres in layer VI of the cortex, and a reduced length of MBP-IR projections extending into

other cortical layers. MBP is translocated from the OL cell body where it is made, up through the OL

processes that extend into the cortex, wrapping and compacting to form myelinated processes

(Pedraza et al., 1997). Thus it is possible that DITPA could be slowing the transport of MBP from

the OL cell body into the OL processes, resulting in a ‘build up’ of MBP closer to the OL body in

cortical layer VI. Although there is no evidence that DITPA directly impacts MBP, DITPA (1ng/mL

or 10ng/mL), it promotes OL differentiation and myelination in human OL co-culture (Lee et al.,

2017).

In the fimbria, DITPA treatment significantly increased the % AC of MBP-IR in both IUGR and

control pups in the present study. The fimbria is situated temporally in the hippocampus, and contains

bundles of afferent and efferent fibres which travel out through the fornix into the rest of the brain.

Thus, in both control and IUGR, DITPA could be having an effect on the processing potential in the

brain of these pups. Future studies should aim to investigate the functional impact of such alterations

due to DITPA, perhaps using motor function testing via the open field test to assess locomotor speed

of rodents (Basso et al., 1995), or the rotor-rod (Deacon, 2013) to assess balance and/or motor

coordination; outcomes can then be correlated with the degree of MBP-IR staining and myelination

present in the cortex and other brain regions, as motor skill learning is correlated with WM structure

(Sampaio-Baptista et al., 2013). There is currently no functional data in humans or animals following

DITPA administration, and this testing would provide useful insight.

PLP-IR

In the present study, IUGR compared to control pups treated with saline showed reduced % AC of

PLP–IR in cortical layer VI, corpus callosum, external capsule and CA1 region of the hippocampus,

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as well as in the fimbria, but no difference in the percentage of cerebral cortex occupied by PLP-IR

projections. These results are consistent with previous studies that report reduced PLP-IR fibres in

the corpus callosum of IUGR rat pups at P14 and P21 (Reid et al., 2012), and in the cerebral cortex

of IUGR guinea pig foetuses (60 dg) compared to controls (Tolcos et al., 2011). In the present study,

in IUGR pups, DITPA compared to saline treatment, significantly decreased the % AC of PLP-IR in

the corpus callosum, and the external capsule. In controls, DITPA compared to saline treatment

decreased the % AC of PLP-IR in the external capsule and the CA1 region of the hippocampus. PLP

constitutes a large proportion of the total myelin proteins, and this decrease could affect the

processing potential of the cerebrum, and as discussed above, motor function testing could be useful

in assessing the implications of this decrease. MBP and PLP are both myelin proteins, and based on

a previous IUGR study in guinea pigs, where both MBP and PLP were reduced (Tolcos et al., 2011),

it was expected that PLP would follow the same trend as MBP in the current study. This is the first

study to examine the effect of DITPA administered to IUGR rodents on PLP levels. The results of

the present study indicate that PLP expression may be more vulnerable to the effects of IUGR than

MBP, with a reduction in the % AC of PLP-IR seen in the external capsule and hippocampus of IUGR

pups, compared to reductions in the % AC of MBP-IR in only the hippocampus.

DITPA treatment in IUGR, and control pups significantly decreased PLP-IR in regions where MBP-

IR was unaffected, or was even elevated, like in cortical layer VI, the corpus callosum, external and

hippocampus (CA1) in controls. PLP is an essential constituent of the myelin sheath, and is

fundamental for myelin compaction and wrapping, therefore decreased coverage could signify

impaired construction of the myelin sheath, or wrapping around axons in these brain regions. It would

be useful to examine myelinated axons in these areas of the brain using transmission electron

microscopy in order to better determine if DITPA is at all negatively affecting myelin compaction or

wrapping.

In the present study, both MBP- and PLP-IR were examined as they have different yet essential roles

in formation of the myelin sheath – PLP is essential for compaction and wrapping of the myelin

strand, while MBP is fundamental for adhesion of myelin layers and signalling to OLs (Boggs, 2006,

Dyer et al., 1994). Therefore assessing MBP- and PLP-IR in the brain provides a more complete

representation of myelination than simply examining MBP or PLP alone. Within the myelin

membrane, MBP is located internally, situated in the cytoplasmic interface, while PLP transverses

the myelin membrane and is exposed on the outer surface (extracellular face) (Baumann and Pham-

Dinh, 2001). PLP is highly hydrophobic and upon synthesis it is transported whole to the plasma

membrane before it reaches the myelin sheath (Baron and Hoekstra, 2010). In contrast MBP is

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hydrophilic in structure and is transported directly, and therefore more quickly, to the myelin sheath

after it is synthesised, albeit in mRNA granule form and not as a complete protein like PLP (Boggs,

2006, Baron and Hoekstra, 2010). The differing polarities and transport time of these myelin proteins,

may underlie the reason why PLP is more impacted by DITPA than MBP, however this theory

requires further investigation.

3.8.2.2 Oligodendrocytes

In the CNS, OLs make myelin and undergo a lineage progression from progenitors to a mature

myelinating phenotype. APC labels only mature myelinating OLs within the brain, while Olig2 labels

the whole OL lineage. Together, these immunostains were used to examine the impact of IUGR and

DITPA treatment on OLs in the WM and GM of the cerebrum. At P14, the areal density of the entire

OL lineage (Olig2-IR; cells/mm2) was reduced in cortical layer VI of IUGR compared to control

pups. This finding differs to our previous study in IUGR rats that found no difference in Olig2-IR

areal density within the corpus callosum at P7, P14 and P35 (Azhan, A., PhD thesis, Monash

University, 2019). It is unclear why this is the case, as the surgical protocol (induction of IUGR) and

species used were identical. However, the two studies were performed across two different institutes,

and the animals were from different breeding facilities, and this may have contributed to the

conflicting results. In the present study the areal density of mature APC-IR OLs, was reduced in

cortical layer VI, the corpus callosum, hippocampus and fimbria of IUGR + saline compared to

control + saline pups, and the percentage of APC-IR mature OLs within the entire population of OLs

was reduced in cortical layer VI and the corpus callosum; reduced density of mature APC-IR OLs in

IUGR is consistent with finding from our previous IUGR rat study (Azhan, A., PhD thesis, Monash

University, 2019). Our finding together with those of the previous study indicates that there are less

mature OLs relative to the entire population of OLs in IUGR, in line with work suggesting that IUGR

delays OL maturation at the pre-myelinating stage (Tolcos et al., 2011). Although there are currently

no clinical studies that have examined the effect of DITPA on OLs in the IUGR brain, an in vitro

study DITPA promotes differentiation and maturation of OL progenitors into mature myelinating

OLs (Lee et al., 2017). However in the present study there was no difference in the number of mature

OLs in comparison to the overall OL population in IUGR pups treated with DITPA. This is an

interesting finding, as DITPA markedly increased MBP in cortical layer VI and fimbria, however did

not appear to alter the number of OLs, indicating that DITPA may be increasing the myelinating

potential of existing mature OLs. Interestingly, when given to control pups, DITPA decreased the

density of Olig2-IR OLs in cortical later VI, as well as mature APC-IR OLs, and decreased the

percentage of mature APC-IR OLs to the overall OL population in cortical later VI and the fimbria.

These results suggest that DITPA being given to controls may be having a detrimental effect on OL

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maturation in the cortex and fimbria, possibly due to an excess of TH (in the form of DITPA) being

given in an otherwise normally functioning system, and thus caution should be taken.

3.8.3 Effects of daily DITPA administration on inflammation in the cerebrum

In the CNS, activation of microglia (microgliosis) and astrocytes (astrogliosis) is the brains response

to injury and is indicative of an inflammatory response (Mayo et al., 2014, Ramlackhansingh et al.,

2011). Both glial cell responses have previously been reported in the brains of IUGR rodents.

Specifically, a trend towards an increase in the number of GFAP-IR astrocytes (Reid et al., 2012) and

a 2.5-fold increase in the number of Iba1-IR microglia (Zanno et al., 2019), has been reported in the

corpus callosum of the postnatal IUGR rat compared to controls. In the brain of IUGR foetal guinea

pigs, a significant increase in GFAP-IR astrocytes and Iba1-IR microglia has been reported in the

cerebral WM (Tolcos et al., 2011) but not in the foetal hippocampus (Cumberland et al., 2017).

Despite these findings, in the present study there was no evidence of microgliosis or astrogliosis in

the IUGR rat brain at P14, in any of the cerebral regions examined. These results were unexpected

based on previous IUGR studies, and at present we have no definitive explanation, however we

speculate that elevated cortisol levels due to an extended injecting regimen may play a role. In cortical

microglial cultures from P1 and P2 rats cortisol blocks microglial activation which is thought to be

an indirect neuroprotective response (Drew and Chavis, 2000). However, whether this is also true for

astrocytes is unknown. In vivo studies show contradictory results. Studies demonstrate that chronic

stress halts astroglial development from precursors (Sabolek et al., 2006), and decreases GFAP

mRNA expression and density of astrocytes following chronic exposure to corticosterone (Nichols et

al., 1990, O'Callaghan et al., 1991) and exposure to early life stress in adult rats (Leventopoulos et

al., 2007). However another study shows a 30% increase in the density of GFAP-IR astrocytes

following a stressor (activity stress) (Lambert et al., 2000). An important finding from the present

study is that DITPA administered to either control or IUGR rat pups, did not lead to cerebral

astrogliosis or microgliosis. This is the first study to examine the impact of DITPA on inflammatory

responses in the postnatal brain, however this requires further investigation at multiple and later

postnatal ages before DITPA can be deemed a safe translational therapy for IUGR babies.

3.8.4 Limitations of the study

This study found that myelination, when assessed by analysing the % AC of MBP-IR, was not

different in cortical layer VI, external capsule and fimbria of IUGR compared to control pups. This

was a surprising result given previous literature (Olivier et al., 2005, Reid et al., 2012, Tolcos et al.,

2011) and it was hypothesised that a stress response to daily injections for an extended period (i.e. 13

days) may underlie the differing results. To support or refute this hypothesis additional “absolute

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control” groups – i.e. control and IUGR pups without intervention (daily handling, weighing,

injecting) –would be necessary. This would allow me to elucidate the effects of IUGR vs. control

without the potential confounding variable of stress. At the beginning of 2020 additional “absolute

control” groups were generated (control: n= 4 litters, IUGR: n= 3 litters). Pups were not handled,

apart from being removed with minimal disturbance when the cage was cleaned once a week. Males

were chosen from this new cohort to match the standard methodology (1 male per litter: n= 4 control,

n= 3 IUGR), and tissue was collected and processed as described in Chapter 2 (Section 2.7). Paraffin

sectioning of this tissue was underway in preparation for MBP-IR staining, when the University was

closed due to the COVID19 pandemic, and as such this task was unable to be completed in time to

include it in this thesis. However the MBP-IR staining of these new cohorts will be analysed and

compared to the experimental groups presented in this thesis, at a later point. Before the COVID-

related lockdown occurred preparing was underway to assess the levels of glucocorticoid receptor

(GR) and mineralocorticoid receptors (MR) in the frozen brain tissue collected, in order to confirm

the presence or absence of a stress response. This was going to be accomplished using quantitative

polymerase chain reaction (qPCR) to measure the gene expression of GR and MR in the hippocampus

of rat pups across experimental groups at P14. Unfortunately saliva was not collected for a cortisol

assay, and no plasma was left following the neonatal plasma assays. Another limitation of this study

was that the immune-staining density and not intensity of both GFAP and Iba1 was measured.

Potential hypertrophy (activation) of astroglia (GFAP-IR) and microglia (Iba1) could have been

indicated by increased intensity of these stains per cell or per area, both of which might have given

an insight into a subtle inflammatory activation. Animal models of IUGR show mixed results

regarding the presence of glial activation. Microglial hypertrophy (Iba1-IR) seen in rats at P3 & P10

(Pham et al., 2015), while an IUGR guinea pig model showed no difference in microglial morphology

at P60 (Tolcos et al., 2015).Gender differences may play a role, with (Fung et al., 2012) demonstrating

that male IUGR rats have a greater density of hippocampal astrocytes than females when compared

to non IUGR counterparts. It is important for future studies to take gender differences into

consideration when examining these glial populations. Researchers are yet to identify whether DITPA

causes any hypertrophy of astroglia and microglia, however this is an extremely important avenue

that future studies should investigate. Lastly, only males were used in this study to maintain

consistency with our previous short-term DITPA study in rats (Azhan, A., PhD thesis, Monash

University, 2019), and because previous studies examining myelination in IUGR rat pups did not

specify sex (Reid et al., 2012, Olivier et al., 2007). However, it should be acknowledged that it is

equally important to examine the effects of both IUGR and DITPA in females; the brains from

females were also collected and processed as outlined in this thesis and these will be studied at a later

point in time.

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3.8.5 Conclusion

This study showed that DITPA administered daily to IUGR rat pups from P1 to P13 may improve

myelination in the cerebrum by P14 as indicated by elevated MBP-IR in cortical layer VI and in the

fimbria, but may also delay migration of myelin proteins into cortical processes and impair expression

of the myelin protein PLP. DITPA appeared to promote the density of Olig2-IR OLs in the corpus

callosum, but there was no difference in the percentage of mature OLs to the overall OL population,

indicating that DITPA does not impact the rate of OL maturation in IUGR, but may inhibit OL

maturation in cortical layer VI and the fimbria, as well as PLP expression when administered to non-

IUGR animals. Importantly, DITPA did not cause injury or inflammation in any cerebral regions in

IUGR rats. The functional consequences of less myelination of cortical processes in response to

DITPA are yet to be determined; this may be transient, however the myelinating potential of DITPA

remains promising. DITPA administration in control animals provided unfavourable outcomes,

reducing PLP, Olig2 and APC expression. Further exploration of DITPA as a neuroprotective utility

should examine the neurodevelopmental effects across multiple species. Please see thesis Chapter 6

(Section 6.5) for further directions, including investigating an alternate route of DITPA

administration and behavioural studies.

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4 Impact of DITPA treatment on

myelination and inflammation

in the neonatal IUGR rat

cerebellum

4.1 Preamble

The overall aim of this thesis is to examine the potential therapeutic benefits of DITPA treatment on

brain development in IUGR. As described previously (Chapter 3), in the cerebrum DITPA treatment

in IUGR pups may improve myelination at P14, evident by elevated MBP-IR staining in the cerebral

cortex compared to saline treated IUGR pups. DITPA did not cause inflammation or disruption to the

cerebrum, but did result in shorter MBP-IR cortical processes, perhaps due to altered transport of

MBP into the processes. The results from Chapter 3 suggest that daily DITPA administration from

P1 to P13 is not having a detrimental effect on the cerebrum of IUGR pups, and DITPA may be

having a beneficial effect on myelination at P14. The next study in this thesis examined the effect of

DITPA administration in another important brain region with known vulnerability to IUGR, the

cerebellum.

4.2 Introduction

The cerebellum is the component of the hindbrain, responsible not only for motor coordination,

balance, equilibrium and muscle tone (Albert et al., 2007), but also cognition and higher order brain

functions (O'Halloran et al., 2012). It is comprised of WM and the highly folded GM of the cerebellar

cortex, and contains approximately half of the total number of neurons in the brain, suggesting it

possesses powerful mechanisms for processing information. The cerebellum increases roughly five-

fold in volume between 24 and 40 weeks post conception (Chang et al., 2000). During this protracted

period of growth and development the cerebellum is highly vulnerable to insults.

It is known that IUGR often results in impaired functional outcomes, with IUGR babies at an

increased risk of neurodevelopmental sequelae (Geva et al., 2006) such as cerebral palsy (McIntyre

et al., 2013). A number of clinical studies have used MRI to investigate the impact of IUGR on the

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brain structure of infants (Padilla et al., 2014, Egana-Ugrinovic et al., 2013, Batalle et al., 2012, De

Bie et al., 2011). Disruptions in regional connectivity, consistent with decreased WM volume have

been found in the cerebella of IUGR infants at 1 year of age (Batalle et al., 2012), and decreased

volumes of cerebellar GM and WM have also been reported in the brains of IUGR children at 12

months, and up to 7 years of age (De Bie et al., 2011, Padilla et al., 2011). These studies suggest that

IUGR negatively impacts the foetal cerebellum, resulting in decreased WM and GM volume. Post-

mortem studies of the human IUGR brain are rare, and there is no current post-mortem literature on

the IUGR cerebellum.

Animal studies examining the effect of IUGR on the cerebellum have shown reduced volumes of the

ML, IGL and cerebellar WM in the guinea pig in response to IUGR, at 60 dg, and at 1 week postnatal

age. Neuronal density was decreased in the IUGR cerebellum, as were the number of Purkinje cells,

the main motor output cell of the cerebellum, at 1 week postnatally (Mallard et al., 2000, Tolcos et

al., 2018). An increase in the volume of the cerebellar EGL, the site of granule cell proliferation is

seen in both the IUGR guinea pig at 60 dg, and the rat at P7 (Tolcos et al., 2018, McDougall et al.,

2017). In the rat, the newborn (P1) IUGR rat cerebellum displays increased rates of neuronal

apoptosis compared to controls (Liu et al., 2011), and there is disorganisation of BG fibres, the

migratory scaffold which granule cells use to migrate to the IGL, at P7 and P35, as well as a 10%

decrease in their linear density at P35 (McDougall et al., 2017). There is currently no treatment for

IUGR-induced cerebellar injury, however given that TH is important for brain development (Bernal,

2007, Escobar et al., 2004), including cerebellar development (Koibuchi, 2008), TH based treatments

should be explored.

In the cerebellum, TH is fundamental for the formation of synapses between cerebellar neurons,

proliferation and migration of cells and branching of Purkinje cell’s dendritic arbours (Nicholson and

Altman, 1972b, Nicholson and Altman, 1972a, Clos et al., 1980, Legrand, 1967). In the IUGR human

(Chan et al., 2014) and rat (Azhan, A., PhD thesis, Monash University, 2019) brain the exclusive TH

transporter MCT8 is reduced, and this may lead to impaired TH signalling and affected myelination

as discussed in Chapter 3. Indeed, myelination is also reduced in the cerebellum, due to disrupted

maturation of oligodendrocytes at the pre-myelinating stage (Clos et al., 1980, Tolcos et al., 2011).

Therefore with reduced MCT8 in the IUGR brain (Aarum et al., 2003, Azhan, 2019, Chan et al.,

2014), treatment with conventional TH may be ineffective. TH analogues such as DITPA could

therefore be a potential therapies to improve myelination as in the IUGR brain, as DITPA does not

require MCT8 for cellular uptake.

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DITPA readily enter brain cells in MCT8 knockout mice, and subsequently correct cerebral

hypothyroidism (Di Cosmo et al., 2009). A previous study by our group found that DITPA

(0.5mg/100g; i.p.) administered from P1 to P6 in IUGR rat pups, a timeframe equivalent the brain

development of an IUGR infant born preterm, promoted myelin recovery, detectable by day 7 in the

external capsule (Azhan, A., unpublished thesis, 2019). Although promising, this data only provides

evidence of the impact of short-term DITPA treatment, and is limited to only examining cortical WM.

It is important to determine whether DITPA can be used as a therapy for the IUGR brain overall, not

just the cerebral WM and GM. Therefore this Chapter specifically aimed to examine the impact of

DITPA treatment on the IUGR cerebellum. In contrast to our previous short-term DITPA study

(Azhan, A., unpublished thesis, 2019), my experimental protocol uses a longer-term treatment

timeframe (DITPA administered from P1 – P13), which is equivalent in human brain development in

a preterm IUGR baby receiving DITPA until term equivalent age.

Therefore this study aimed to: determine whether DITPA administered from P1 to P13 in IUGR

rats (i) impacts cerebellar growth and morphology, (ii) promotes OL maturation, and restore

myelination, and (iii) does not cause cerebellar injury or inflammation. The hypothesis was that

DITPA administration from P1 to 13 will promote and restore OL maturation and myelination within

the cerebellum of IUGR rat pups when assessed at P14, without causing injury or inflammation. This

study will generate complementary information to our previous short-term DITPA administration

study (Azhan, A., unpublished thesis, 2019) and will inform clinical translation of therapy.

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4.3 Methodology

4.3.1 Animals and tissue

The methodology in this Chapter will commence from the point of paraffin tissue sectioning, and will

focus on the cerebellum. For comprehensive methodology regarding animal welfare, species, surgical

procedure, drug treatment, post-mortem and tissue collection, please refer to Chapter 2 (Sections 2.1

to 2.8).

4.3.2 Paraffin sectioning of the cerebellum

Paraffin embedded P14 rat cerebellums were sectioned sagittally at 8µm using a rotary microtome

(Jung Biocut 2035, Geprufte Sicherhiet, Germany). A total of 40 sections were cut from each

cerebellum (n= 8 control + saline, n= 7 control + DITPA, n= 8 BUVL + saline, n= 8 BUVL + DITPA;

1,240 sections in total) and sections were placed in a warm water bath (~ 40°C) prior to being mounted

flat onto slides (Superfrost plus +, Menzel Glaser, Germany). Two sections per animal were placed

onto slides and each section was separated by 80µm (8µm x 10; Figure 4.2 A). All tissue-mounted

slides were dried overnight at 30°C prior to histological and immunohistochemical staining.

4.3.3 Assessment of the cerebellar structure

H&E staining was performed using a standard procedure (Chapter 2 section 2.9.1). H&E stained

sections (2 slides/animal = 4 sections/animal; 124 sections total) were digitally scanned using

Olympus slide scanning software at 20x magnification, and using a constant light setting (Olympus

VS120-S5 scanner, Olympus, Vic, Australia); the digital images were visualised using Fiji (ImageJ,

Version 2.0, National Institute of Health, Maryland, USA). The area (mm2) of the IGL (containing

predominantly post-mitotic granule cells), ML (containing Purkinje cell dendrites and parallel fibres

from granule cells), GM (containing nerve cell bodies, dendrites and axon terminals) and the WM

(containing myelinated and unmyelinated afferent and efferent fibres) as well as the total cerebellar

cross-sectional area was traced using the Fiji trace function (Figure 4.1). Areas of the ML, IGL, GM

and WM were expressed as a ratio of the total cerebellar cross-sectional area for each section, and

means calculated per animal and per group.

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The width of the ML was assessed at P14 using x20 magnification. The Fiji software ‘line’ tool and

‘measure’ function were used to measure the width of the ML (µm) in 20 random places in each of

the early and late developing lobules (X and VIII; 20 measurements/lobule). The average width of all

of the ML measurements was averaged per lobule, per section and per animal. Differences in

cerebellar ML width following IUGR would be indicative of disturbance in normal development and

signalling, as the ML contains the dendritic trees of Purkinje cells, as well as a large population of

interneurons.

4.3.4 Immunohistochemical staining of the cerebellum

Immunohistochemistry was performed using the standard protocol outlined in Chapter 2 (Section

2.10). A number of trials were performed for each antibody to ensure optimal staining quality prior

to conducting the procedure on the actual tissue. For a full outline of final conditions used for each

immunostain performed in the cerebellum, please refer to Table 4.1. To avoid procedural variation,

and ensure uniform conditions for subsequent analysis, sections from all experimental groups were

stained simultaneously for each antibody/procedure. Sections were counterstained with

Haematoxylin, with the exception of MBP and GFAP. In addition, positive controls (foetal sheep

cerebellum) and negative controls (omission of primary antibody) were performed for each antibody.

Figure 4. 1 H&E stained sagittal section of P14 rat cerebellum at the level of the vermis. The total cross-

sectional area of the cerebellum (yellow trace), molecular layer (ML; between yellow and green trace),

internal granular layer (between green and blue trace), white matter (blue trace) and grey matter (area of ML

+ area of internal granular layer) were traced and measured. ML width measurements were conducted in

early (I or X) and late developing lobules (VII or VIII). Scale bar = 1mm.

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4.3.5 Immunohistochemical analysis of the cerebellum

Prior to quantitative assessment, an independent researcher coded sections, to ensure that analyses

were performed blinded to experimental group. Analyses were conducted on 4 sections per animal,

using Fiji software (as for H&E analysis, Section 4.3.3). Sections were separated by 80m, and 2

slides per animal were used for each stain (see figure 4.2 for cutting and staining progression).

Measurements were made in both an early (I or X) and a late developing lobule (VII or VIII) for each

analysis. The lobules were selected to assess the full developmental range of the cerebellum.

Measurements were averaged per lobule for the section, and per animal (average of 4 sections/animal)

and unless there were differences between early and late developing lobules, data were combined and

averaged and presented as simply ‘lobules’.

Figure 4. 2 Sequence of tissue sectioning and staining (A) A total of 40 sections were cut from each tissue

block; 8µm apart. (B) 2 slides, and hence 4 sections per animal were used for each immunostain (control +

saline n= 8, IUGR + saline = 8, control + DITPA = 7, IUGR + DITPA = 8; total 31 animals; 2 slides/ animal

= 62 slides for each stain). GFAP = glial fibrillary acidic protein, H&E = Haemotoxylin & Eosin, IBA1 =

Ionised calcium binding adaptor molecule – 1, MBP = myelin basic protein, Olig2 = oligodendrocyte

transcription factor 2.

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4.3.5.1 Myelin basic protein (MBP)-immunoreactivity (IR)

Areal coverage (% AC) of MBP-IR fibres was assessed in the lobular and deep WM (DWM) of each

section. Four images were taken from the DWM (200 x 200m = 20,000m2), and three images were

taken from the lobular WM (100 x 100m = 10,000m2) at 20x magnification using Photoshop

software ‘set measurement’ square crop tool (Photoshop CC, v 19.0, Adobe Systems Incorporated).

Fiji threshold analysis (ImageJ, Version 2.0, National Institute of Health, Maryland, USA) was

carried out on each image, using a constant threshold for all tissue. Data were averaged per region

(DWM or lobule), per section, and then per animal. Threshold analysis measures differences in

staining intensity, and a threshold of 150 (reflecting dark staining) was used to determine the

proportion (%) of WM or DWM occupied by MBP-IR. If there were no differences in % AC between

early and late developing lobules, they were averaged and presented combined as ‘lobules’.

4.3.5.2 Oligodendrocyte transcription factor 2 (Olig2)–IR

Analysis of Olig2-IR cells in the DWM and lobular WM of each section was performed using the

Adobe Photoshop Software to extract regions of interest (ROIs) using the same dimensions and

protocol as described in Section 4.3.5.1 for MBP-IR; Fiji software (ImageJ, Version 2.0, National

Institute of Health, Maryland, USA) was used to count positively stained cell bodies. Olig2-IR cells

were counted in four 200 x 200 µm ROIs (field size = 20,000 µm2) positioned randomly in the DWM

of each section and three ROIs positioned in each of the early and late developing lobules (100 x 100

µm; field size = 10,000 µm2) (Figure 4.3). Olig2-IR cells were identified as having a defined cell

body with obvious dark staining. The average number of Olig2-IR cells in each ROI was counted and

divided by the ROI area to give an areal density of Olig2-IR OLs (cells/mm2). A mean density was

calculated for the DWM and lobular WM of each section and then per animal. If there were no

differences in cell density between early and late developing lobules, they were averaged and

presented combined as ‘lobules’.

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4.3.5.3 Ionized calcium-binding adaptor molecule 1 (Iba1) – IR

Areal density (cells/mm2) of Iba1-IR microglia was assessed in the cerebellar DWM and in the lobule

WM (early or late developing lobules), using the same methodology described for Olig2-IR OL

analysis (Section 4.3.5.2).

4.3.5.4 Glial fibrillary acidic protein (GFAP) - IR

Analysis of the % AC of GFAP- IR fibres was carried out in fields of view within the cerebellar

DWM and lobular WM (early or late developing lobules), in the same way as was described for MBP-

IR, with the exception of a different staining threshold (Section 4.3.5.1). Qualitative analyses of

differences in BG morphology, as well as an assessment of the linear density of BG fibres (fibres/mm)

were carried out in each of the early and late developing lobules per tissue section. Using the Fiji line

measure tool (ImageJ, Version 2.0, and USA), 10x 100µm horizontal lines were positioned randomly

in the middle of the ML of each lobule at 20x magnification, so as to intersect the BG fibres. The

number of fibres crossing each line was counted and divided by the length of the line and the average

Figure 4.3 Olig2-immunostained sagittal section of P14 rat cerebellum at the level of the vermis

(counterstained with Haematoxylin). Quantitative analysis of Olig2-IR cells (arrow in inset) was performed

in the cerebellar white matter, with four regions of interest (ROIs) randomly positioned in the deep white matter

(DWM), and three ROIs placed in the WM of an early developing lobule (Lobule I) and a late developing

lobule (Lobule VIII). Scale bar = 1mm. 20x = 20 x magnification.

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number of counts was calculated per lobule. If there were no differences in linear density of BG

between early and late developing lobules, these were averaged and presented combined as ‘lobules’.

4.3.5.5 Calbindin-IR Purkinje cells

Calbindin is a calcium-binding protein, which is specific to Purkinje cells in the cerebellum. Purkinje

cells are the main output cell of the cerebellum and therefore play an important role in cerebellar

function. To observe any alterations in Purkinje cell development, the somal area of 20 calbindin-IR

Purkinje cells were randomly selected from each of the early and late developing lobules, and the

somal area measured using the Fiji trace and measure functions (40 cells/cerebellum section; 4

sections = 160 cells/animal). Only calbindin-IR Purkinje cells with a visible and clearly defined

nucleus were selected, to allow for an accurate measurement of soma circumference. Data was

expressed as average somal area (µm2), and means taken per section and per animal.

Linear density analyses of Purkinje cells were carried out in each of the early and late developing

lobules per section. Using the Fiji line measure tool, 5 x 100µm horizontal lines were positioned in

the middle of the Purkinje cell layer of each lobule at 20x magnification, so as to intersect the cell

somas. The number of cell somas crossing each line was counted, and divided by the line length to

give the number of Purkinje cells/mm. To calculate the areal density of Purkinje cells, the diameter

was taken from each Purkinje cell which intersected the measurement line, and with this, the areal

density of Purkinje cells was calculated by dividing the number of Purkinje cells/mm by the tissue

section thickness (= mean diameter of Purkinje cell + section thickness (8µm) (McDougall et al.,

2017a).

4.3.6 Statistical analysis

All statistical analyses were performed using the Graphpad Prism statistical software (Graphpad

Software, Version 8.0, CA, USA). Prior to analyses, outliers were removed using the Grubb’s test to

determine a significant outlier (p < 0.05). Data were checked for normal distribution, and if found to

be not normally distributed, the appropriate non-parametric test used (Mann-Whitney U test, instead

of unpaired t-test; Friedman test instead of two-way ANOVA). A two-way ANOVA was used, with

a post-hoc pairwise comparison to identify between group differences, and a Bonferroni correction

was used for multiple post-hoc comparisons between experimental groups. Data were considered

significant if p < 0.05. All main effects data are presented in text as ratio of residual variances (F) and

statistical significance (p-value). All post-hoc data are presented as [mean comparative difference

(95% confidence interval); and p-value], and graphs are represented as Mean standard error of the

mean (SEM). Unless there were differences between early and late developing lobules, data will be

Chapter 4

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presented combined as ‘lobules’. All two-way ANOVA results are tabulated in Appendix 4 as Mean

SEM.

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Table 4. 1 Immunohistochemistry - optimised antigen retrieval, blocking protocol and antibody concentrations in the cerebellum.

Abbreviations: Ab, Antibody; APC, adenomatous polyposis coli; BSA, bovine serum albumin; cat, catalogue number; DAB, diaminobenzidine; GFAP, glial fibrillary

acidic protein; H2O2, hydrogen peroxide; Iba1, ionized calcium-binding adaptor molecule 1; M, molar; MBP, myelin basic protein; Olig2, oligodendrocyte transcription

factor 2; PBS, phosphate buffered saline; PLP, myelin proteolipid protein.

Protein Staining for Antigen Retrieval & block 1°Ab (source); conc. 2°Ab (source); conc. Complex binding

to secondary and

DAB

[Complex][DAB

]

MBP Myelin in the

CNS




min; 4% BSA in PBS

Rat anti-MBP,

(Millipore cat:

MAB386); 1:250

Biotinylated rabbit anti-

rat,

(Millipore); 1:200

Avidin-biotin

complex

DAB (in stable

peroxide buffer)

50μl A + 50μl B

in 10 ml PBS;

1 DAB tablet in

10ml dH2O +

3μl H2O2

Olig2 Entire OL lineage Heat in 0.01M citrate buffer (pH



min;

4% BSA in PBS

Rabbit anti-Olig2,

(Millipore cat:

MABN50); 1:500


rabbit,

(Vector cat:

VEBA1000); 1:200

GFAP BG & astrocytes 1:500 Proteinase K in 37°C for 30

min; 4% BSA in PBS

Rabbit anti-GFAP,

(DAKO cat: 0334):

1:500


rabbit,

(Vector cat:

VEBA1000); 1:200

Iba1 Microglia &

macrophages in

CNS




min;

4% BSA in PBS

Rabbit anti-Iba-1,

(Wako cat: 019-19741);

1: 1000


rabbit,

(Vector cat:

VEBA1000); 1:200

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4.4 Results

4.4.1 Morphology of the cerebellum

There was no qualitative evidence of lesions, infarcts or morphological abnormalities in H&E-stained

cerebellar sections from IUGR or control pups treated with saline or DITPA.

4.4.1.1 Cerebellar layer areas

Two-way ANOVA results showed no interaction between group and treatment on total cerebellar

cross-sectional area (TCA), or area of the ML, IGL, WM or GM (all mm2) (Figure 4.4). There was a

main effect of group on TCA, and area of the ML and IGL (TCA: F1/24 = 10.34, p = 0.0037; ML: F1/21

= 10.23, p = 0.004; IGL: F1/19 = 7.91, p = 0.01), and treatment on area of GM and WM (WM: F1/20 =

7.53, p = 0.01; GM: F1/18 = 3.18, p = 0.09). Post-hoc analysis revealed that there was no difference in

the TCA (Figure 4.4 A), or thicknesses of the ML, IGL, WM or GM (Figure 4.4 B - E) in control

pups compared to IUGR pups, and when treated with DITPA or saline.

Ratios of the cerebellar layer areas to TCA were determined (cerebellar layer area/TCA = %) to adjust

for the effect of confounding variables such as difference in expansion of the tissue in the water bath

(as described in Chapter 3, Section 3.3). Two-way ANOVA results showed no interaction between

group and treatment on ML:TCA, IGL:TCA, WM:TCA or GM:TCA (Figure 4.4 F-I), and no main

effects of group or treatment. Post-hoc analysis revealed no difference in ML:TCA, IGL:TCA,

WM:TCA or GM:TCA (Figure 4.4 F - I) in control pups compared to IUGR pups, treated with DITPA

or saline.

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4.4.1.2 Cerebellar molecular layer width

Two-way ANOVA results showed no interaction between group and treatment on width of the ML

(mm) in the cerebellar lobules (F1/50 = 0.68, p = 0.41), however there was a main effect of both group

and treatment on ML width (group: F1/50 = 4.11, p = 0.05; treatment: F1/50 = 4.28, p = 0.042). Post-

hoc analysis revealed that there was no difference in ML width in the cerebellar lobules of control

pups compared to IUGR pups, and when treated with DITPA or saline (Figure 4.5).

Figure 4.4 Total cerebellar cross-sectional-area (A), layer widths (B – E) and layer width-to- total cross-

sectional-area (TCA; F – I) in control and IUGR pups treated with DITPA or saline at P14. Data analysed

using a two-way ANOVA with Bonferroni correction. Values presented as Mean ± SEM. Pup numbers: control

+ saline: n = 8; control + DITPA: n = 7; IUGR + saline: n = 8; IUGR + DITPA: n = 8. All animals are male.

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4.4.2 Immunohistochemical assessment of the cerebellum

4.4.2.1 MBP-IR

Two-way ANOVA results showed no interaction between group and treatment on the % AC of MBP-

IR in the deep WM (DWM; F1/25 = 0.02, p = 0.88), or cerebellar lobules (F1/26 = 0.47, p = 0.50). There

was no main effect of group or treatment. Post-hoc analysis revealed no difference in the % AC MBP-

IR in the DWM (Figure 4.6 A) or lobule WM (Figure 4.6 B) between control and IUGR pups, and

when treated with DITPA or saline.

4.4.2.2 Olig2-IR

Two-way ANOVA results revealed no interaction between group and treatment on the cell density of

Olig-2-IR OLs (cells/mm2) in the DWM or lobule WM (DWM: F1/24 = 0.88, p > 0.36; Lobules: F1/26

= 1.80, p = 0.19). There was no main effect of group or treatment. Post-hoc analysis showed no

difference in Olig2-IR cell density in the cerebellar DWM (Figure 4.6 C) or lobule WM (Figure 4.6

D) between control and IUGR pups, and when treated with DITPA or saline.

Figure 4.5 Width of the molecular layer in control and IUGR pups treated with DITPA or saline at P14.

Data analysed using a two-way ANOVA with Bonferroni correction. Values presented as Mean ± SEM. Pup

numbers: control + saline: n = 8; control + DITPA: n = 7; IUGR + saline: n = 8; IUGR + DITPA: n = 8. All

animals are male.

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Figure 4.6 Area coverage of MBP-IR in the cerebellar deep white matter (A) and lobule white matter

(B), and density of Olig2-IR oligodendrocytes in the deep white matter (C) and lobule white matter (D)

in control and IUGR pups treated with DITPA or saline at P14. Representative photomicrographs of MBP-

IR in the DWM of a control + saline (E), control + DITPA (F), IUGR + saline (G) and IUGR + DITPA (H)

pup show no difference in the % AC of MBP-IR between groups. Data analysed using a two-way ANOVA

with Bonferroni correction. Values presented as Mean ± SEM. Pup numbers: control + saline: n = 8; control +

DITPA: n = 7; IUGR + saline: n = 8; IUGR + DITPA: n = 8. All animals are male.

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4.4.2.3 Iba1-IR

Two-way ANOVA results showed an interaction between group and treatment on the density of Iba1-

IR microglia (cells/mm2) in late developing lobules (F1/24 = 6.91, p = 0.01). There was a main effect

of group on Iba1-IR microglia density in late developing lobules (F1/24 = 4.39, p = 0.05), but not in

the early lobules. Post-hoc analysis showed that there was no difference in density of Iba1-IR

microglia in the DWM (Figure 4.7 A) or lobule WM (Figure 4.7 B, C) of IUGR + pups compared to

control + saline, but there was higher density of Iba1-IR microglia in the late developing lobules of

IUGR + DITPA compared to IUGR + saline pups [551.3 (86.17, 921.7); p = 0.008; Figure 4.7 C],

and in IUGR + DITPA when compared to control + DITPA pups [503.9 (86.2, 921.7); p = 0.01;

Figure 4.7 C].

Figure 4.7 Cell density of Iba1-IR microglia in the cerebellar deep white matter (A) and lobule white

matter (B, C) in control and IUGR pups treated with DITPA or saline at P14. Representative

photomicrographs of Iba1-IR microglia in the WM of a late developing lobule in a control + saline (E),

control + DITPA (F), IUGR + saline (G) and IUGR + DITPA (H) pup shows an increase in Iba1-IR microglial

density in IUGR + DITPA pup compared to IUGR + saline and control + DITPA. Data analysed using a two-

way ANOVA with Bonferroni correction. * p < 0.05, ** p < 0.01. Values presented as Mean ± SEM. Pup

numbers: control + saline: n = 8; control + DITPA: n = 7; IUGR + saline: n = 8; IUGR + DITPA: n = 8. All

animals are male.

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4.4.2.4 GFAP-IR Bergmann glia (BG)

Two-way ANOVA results showed no interaction between group and treatment on linear density of

BG cells (cells/mm) in early and late developing cerebellar lobules (Early: F1/27 = 6.6, p = 0.016; Late:

F1/27 = 12.13, p = 0.0017). There was a main effect of group on BG linear density in early and late

lobules (early: F1/27 = 27.38, p < 0.0001; late: F1/27 = 61.81, p < 0.0001). Post-hoc analysis revealed

that there was an increased density of BG fibres in IUGR + saline pups compared to control + saline

pups in the late developing lobules [20.16 (1.95, 38.36); p = 0.024; Figure 4.8 B], and there was a

decrease in the density of BG fibres in control + DITPA pups compared to control + saline [-24.08 (-

42.93, -5.24); p = 0.007; Figure 4.8 B], but no difference in early lobules (Figure 4.8 A). There was

an increased density of BG fibres in IUGR + DITPA pups compared to control + DITPA pups in both

the early and late developing lobules [early: -35 (-54.52, -17.00); p < 0.0001; late: -52.21 (-71.06, -

33.36); p < 0.0001; Figure 4.8 A, B].

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4.4.2.5 Area coverage of astrocytes

Two-way ANOVA results showed no interaction between group and treatment on the % AC of

GFAP-IR astrocytes in the DWM, or lobule WM (DWM: F1/27 = 0.12, p > 0.73; Lobules: F1/56 = 1.13,

p = 0.29). There was only a main effect of group in the lobules (F1/56 = 9.18, p = 0.04). Post-hoc

analysis showed no difference in the % AC of GFAP-IR astrocytes in the DWM or lobule WM of

IUGR + saline pups compared to control + saline, or between IUGR + DITPA and IUGR + saline

pups.

Figure 4.8 Linear density of GFAP-IR Bergmann glia (BG) in the early (A) and late (B) developing

cerebellar lobules in control and IUGR pups treated with DITPA or saline. Representative

photomicrographs of GFAP-IR BG fibres in the ML of a late developing lobule in a control + saline (C), control

+ DITPA (D), IUGR + saline (E) and IUGR + DITPA (F) pup shows an increase in BG linear density in IUGR

+ saline pups compared to control + saline, and decreased linear density in control + DITPA pup compared to

control + saline. Data analysed using a two-way ANOVA with Bonferroni correction. * p < 0.05, ** p < 0.01, **** p < 0.0001. Values presented as Mean ± SEM. Pup numbers: control + saline: n = 8; control + DITPA: n

= 7; IUGR + saline: n = 8; IUGR + DITPA: n = 8. All animals are male.

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There was an increase in % AC of GFAP-IR astrocytes in IUGR + DITPA pups compared to control

+ DITPA pups when data from the lobules were combined [-43.98 (-57.07, -30.90); p < 0.0001; Figure

4.9 B], but not when analysed separately (Figure 4.9 C, D).

4.4.2.6 Calbindin-IR

4.4.2.7 Purkinje cell somal area

Two-way ANOVA results showed no interaction between group and treatment on the somal area

(mm2) of calbindin-IR Purkinje cells, in cerebellar lobules (F1/53 = 0.13, p = 0.72), and no main effects

of group or treatment. Post-hoc results showed no difference between calbindin-IR Purkinje cell

somal size between control and IUGR pups, and when treated with DITPA or saline (Figure 4.10 A).

Figure 4.9 Area coverage of GFAP-IR astrocytes in the cerebellar deep white matter (A), early and late

lobule white matter combined (B), as well as early lobules (C) and late lobules (D) separately, in control

and IUGR pups treated with DITPA or saline at P14. Data analysed using a two-way ANOVA with

Bonferroni correction. * p < 0.05. Values presented as Mean ± SEM. Pup numbers: control + saline: n = 8;

control + DITPA: n = 7; IUGR + saline: n = 8; IUGR + DITPA: n = 8. All animals are male.

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4.4.2.8 Purkinje cell areal density

Two-way ANOVA results showed no interaction between group and treatment on Purkinje cell areal

density (cells/mm2) in the cerebellar lobules (F1, 25 = 0.041, p = 0.84). There was a main effect of

treatment on areal density of Purkinje cells in the early developing lobules (F1, 27 = 9.43, p = 0.005).

Post-hoc analysis showed that there was no difference in calbindin-IR Purkinje cell areal density

between control and IUGR pups, and when treated with DITPA or saline (Figure 4.9 B).

4.4.2.9 Purkinje cell linear density

Two-way ANOVA results showed no interaction between group and treatment on linear density of

Purkinje cells (cells/mm) in cerebellar lobules at P14 (F1/25 = 0.02, p = 0.88). There was a main effect

of treatment on linear density of Purkinje cells in the early developing lobules (F1/27 = 25.29, p <

0.001). Post-hoc analysis showed an increase in the linear density of Purkinje cells in the early

developing lobules of IUGR + DITPA pups compared to IUGR + saline [9.062 (2.13, 16.00); p =

0.006; Figure 4.10 D], and in control + DITPA pups compared to control + saline. There was no

difference between IUGR + saline pups and control + saline [8.57 (1.39, 15.75); p = 0.01; Figure 4.10

D].

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Figure 4.10 Somal area (A), areal density (B) and linear density of calbindin-IR Purkinje cells in

cerebellar lobules combined (C), and in early (D) and late lobules (E) separately, in control and IUGR

pups treated with DITPA or saline at P14. Representative photomicrographs of calbindin-IR Purkinje cells

in an early developing cerebellar lobule in a control + saline (F), control + DITPA (G), IUGR + saline (H) and

IUGR + DITPA (I) pup shows an increased linear density of calbindin-IR Purkinje cells in IUGR + DITPA

pup compared to IUGR + saline and control + DITPA pups compared to control + saline. Data analysed using

a two-way ANOVA with Bonferroni correction. * p < 0.05, ** p < 0.01. Values presented as Mean ± SEM.

Pup numbers: control + saline: n = 8; control + DITPA: n = 7; IUGR + saline: n = 8; IUGR + DITPA: n = 8.

All animals are male.

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4.5 Discussion

4.5.1 Overview

This is the first study to examine the neurostructure of the neonatal cerebellum in response to daily,

longer-term DITPA administration (P1 to 13) in an IUGR rodent model. The key findings are:

In all experimental groups there was no difference in the presence of lesions, haemorrhages or infarcts

between IUGR or control pups given DITPA or saline when assessed qualitatively, and no alteration

to total cerebellar cross-sectional area, or cerebellar layer areas.

In IUGR + saline compared to control + saline pups there was significantly (i) increased linear density

of BG fibres (GFAP-IR) in the late developing cerebellar lobules of IUGR pups (VII or VIII;

cells/mm), and (ii) no difference in the % AC of MBP-IR, or GFAP-IR astrocytes, or in the areal

density of Iba1-IR microglia, or Olig2-IR OLs (cells/mm2) in the DWM or WM of the early or late

developing lobules. These results signified that IUGR did not injure the developing cerebellum.

In IUGR + DITPA compared to IUGR + saline pups there was a significant (i) increase in the areal

density of Iba1-IR microglia in the late developing cerebellar lobules, and (ii) linear density of

Purkinje cells (cells/mm) in the ML of early developing cerebellar lobules.

In control + DITPA compared to control + saline pups there was a significant (i) decrease in the linear

density of GFAP-IR BG fibres in the late developing cerebellar lobules, and (ii) an increase in the

linear density of Purkinje cells in the early developing lobules.

4.5.2 Effect of daily DITPA administration on cerebellar structure

The presence of IUGR did not alter cerebellar morphology, or result in overt injury (lesions,

haemorrhages or infarcts) when assessed qualitatively using H&E staining. There was also no

difference in somal area (mm2), linear density (cells/mm) or areal density (cells/mm2) of Purkinje

cells in IUGR + saline pups compared to control + saline pups. A previous study in IUGR guinea pigs

found that somal area of Purkinje cells was reduced at 60 dg (Tolcos et al., 2018), however differences

in results between this study and the current study are likely due to differences in species used and

therefore the timing of IUGR onset.

WM volumes in the cerebellum of IUGR children assessed clinically using MRI at 1 year of age

(Batalle et al., 2012, Padilla et al., 2011) and at 4 to 7 years of age (De Bie et al., 2011). Although

these findings differ to the findings in the present study it is important to note that these clinical

findings were made at much later developmental time points than the current study, and with

unknown timing of IUGR onset. In animal models, the effect of IUGR on cerebellar morphology

varies between species, and also with timing of both IUGR onset (mid- vs. late gestation), and age of

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analysis (foetal vs. postnatal). The results of the present study are consistent with those of McDougall

and colleagues (McDougall et al., 2017b) who showed no difference in area of the IGL at P7, or the

TCA and areas of the ML, IGL, WM, and ratio of these layers to TCA at P35 in the IUGR rat induced

via late gestation BUVL. The findings of present study are also consistent with those of Tolcos and

colleagues (Tolcos et al., 2018) who showed no difference to TCA or area of the IGL in IUGR

compared to control pups using a guinea pig model of IUGR (unilateral uterine vessel ligation at mid-

gestation). In contrast to the present study, there is a decrease in IGL area in IUGR guinea pigs at 60

dg, and at 1 week postnatal age, along with decreased volume of WM and the ML (Tolcos et al.,

2018); (Mallard et al., 2000).

4.5.3 Effect of daily DITPA administration on white matter development

MBP labels mature myelinated fibres within the brain, while Olig2 labels the whole OL cell lineage;

together, these stains were used to determine the overall effect of DITPA treatment on cerebellar

WM. There was no difference in area coverage of MBP-IR, or Olig2-IR OL density in the cerebella

of IUGR + saline pups compared to control + saline pups. These results are supported by the lack of

difference in total cerebellar WM area in IUGR pups compared to controls (both given saline), when

assessed using H&E (Section 4.3.3). This was a surprising finding, given it is widely reported in

rodent models that IUGR reduces both myelin and OL density in the brain (Tolcos et al., 2011, Reid

et al., 2012, Olivier et al., 2007, Olivier et al., 2005). These previous studies however, were

predominantly conducted in cortical WM, and differences between the cortex and cerebellum must

therefore be taken into account when comparing data. In the foetal guinea pig cerebellum area

coverage of MBP-IR is reduced and area density of Olig2-IR OLs is increased, however both

normalised to control levels by 1 week postnatal age (Tolcos et al., 2011). This raises the question of

whether any differences in myelination in the cerebellum were ‘corrected’ by P14 in the present study.

It is still not clear why a reduction in the % AC of MBP-IR was not seen in the cerebella of IUGR

pups in the present study at P14, however the stress of daily handling and injecting of pups (with

either saline or DITPA) for an extended period of time impacts myelination in control and IUGR pups

alike. As previously discussed (Chapter 3, Section 3.8.2), cortisol is released during a stress response

and this delays myelination of major WM tracts in the brain (Bohn and Friedrich, 1982, Antonow-

Schlorke et al., 2009, Shields et al., 2012). In the present study pups underwent daily physical

handling (e.g. weighing, marking), as well as the physical and psychological stress of intraperitoneal

injections. The surviving OLs in the cerebella of IUGR pups may be more resilient to the effects of

cortisol than controls, having developed under the stress of IUGR conditions (Back et al., 2002, Back

et al., 2001). As discussed previously in Chapter 3, if this were the case, myelination would more

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greatly impaired in control pups compared to IUGR pups in response to stress, thus lowering MBP-

IR to equal that seen in IUGR pups. Future studies should aim to investigate the density and re-

myelinating potential of OLs in the IUGR cerebellum, with and without exposure to daily handling

and compare this to OLs in the non-IUGR rat brain in order to answer this question.

DITPA treatment did not alter MBP-IR levels in the cerebellar DWM, or WM of the early (I or X) or

late (VII or VIII) developing lobules. Currently no clinical or preclinical studies to date have

examined the effects of DITPA treatment on the cerebellum, however DITPA given daily from P1 to

P6 had a restorative effect on myelin in the IUGR rat cerebrum at P7 (Azhan, A., unpublished thesis),

and therefore a similar pro-myelinating effect was expected in the cerebellum. The lack of difference

in MBP-IR coverage in the cerebellum between control and IUGR rats in the present study hinders a

valid assessment of DITPA’s effects on the cerebellar WM. The extended duration of DITPA

administration in the present study (P1 to P13) compared to our group’s previous study (P1 – P6)

appears to be less effective at promoting myelination in IUGR, perhaps due to the stress associated

with an extended injecting regimen. It is impossible to completely eliminate handling stress in pups,

however an alternate route of DITPA administration should be investigated, including oral

administration and nanoemulsions (Anton et al., 2008, Comfort et al., 2015) in order to minimise a

possible stress response to enable correct assessment of DITPA’s effects on myelination.

4.5.4 Effect of DITPA administration on inflammation in the cerebellum.

4.5.4.1 Microglia

IUGR did not affect the density of microglia (Iba1-IR) in the cerebellar DWM or lobular WM. DITPA

administration, on the other hand significantly increased the density of Iba1-IR microglia in the late

developing cerebellar lobules of IUGR pups compared to saline, but not in the early developing

lobules or DWM. Microglial activation in the CNS is typically indicative of an inflammatory response

but are also important for normal brain development (Harry, 2012). Reactive (amoeboid) microglia

are characterised by retracted processes and are classically associated with an inflammatory response

to infection or injury (Graeber et al., 2011), while ramified (resting) microglia are characterised by

long extended processes, and are crucial for neuronal health, as well as clearing dead cells, and even

pruning and remodelling neuronal processes in the foetal brain (Schafer et al., 2012, Walker et al.,

2014). In the present study the thickness of tissue sections (8µm), prevented the definitive

determination of reactive (amoeboid) versus resting (ramified) microglia, as microglial processes

extend in all directions over many microns. Without knowing the state of the microglia in the late

cerebellar lobules, there is insufficient information to comment on the nature of the increase in

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microglial density observed in response to DITPA in the present study. Future studies that observe a

microglial response to DITPA, should therefore investigate microglial morphology using thicker

tissue sections to image the full depth of the section.

There is limited literature surrounding differential vulnerability of cerebellar lobules, however one

study showed that late developing cerebellar lobules in the 10 day old rat (especially VI – VIII) appear

to be most sensitive to chemotherapy (injections of Cisplatin), as cellular differentiation is still

occurring (Pisu et al., 2003) raising the question of whether late developing cerebellar lobules are

more vulnerable to drug treatment compared to early lobules. Future studies should aim to more

closely investigate the differences in reactivity between cerebellar lobules in the presence of DITPA

treatment.

4.5.4.2 Bergmann glial fibres

The ML of the cerebellum contains BG fibres as well as dendritic arbours of Purkinje cells. BG fibres

act as a scaffold along which granule cells migrate from their outer proliferative zone in the EGL to

their final destination in the IGL. Quantitative analysis of GFAP-IR in the ML showed that the linear

density of BG was significantly increased in the late developing cerebellar lobules of IUGR + saline

pups compared to control + saline pups at P14, but not in the early developing lobules. This finding

is somewhat different to a previous study that reported a 10% decrease in linear density of BG in the

cerebellum of IUGR rats at P7 and P35 (McDougall et al., 2017b). DITPA administration had no

effect when given to IUGR pups, but did reduce BG linear density in the late developing cerebellar

lobules when given to control pups compared to saline. Given that BG fibres are a target of TH during

cerebellar maturation (Fauquier et al., 2014), this result could be the effect of DITPA (or excess TH)

being administered in an otherwise functioning system. Fewer BG fibres may indicate disruption to

neuronal migration in the cerebellum. It is not clear why this reduction was only seen in late

developing lobules of control pups and requires further investigation.

4.5.4.3 Astrocytes

Astrogliosis occurs when astrocytes increase in size (hypertrophy) and number (hyperplasia), and is

a response of the brain to injuries. In the present study IUGR alone did not affect areal coverage of

GFAP-IR astrocytes in the DWM or WM of the lobules. In foetal and neonatal rat, guinea pig and

ovine models, astrocyte hypertrophy and hyperplasia occurs in the IUGR brain, compared to controls

(Nitsos and Rees, 1990, Olivier et al., 2007, Tolcos et al., 2011, Rees et al., 1998), however these

studies did not focus on the cerebellum. DITPA treatment, compared to saline had no effect on

astrocyte areal coverage in the DWM or WM of the lobules of control or IUGR pups. However when

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data from lobules were combined there was a significant increase in the areal density of astrocytes in

IUGR + DITPA compared to control pups + DITPA pups, indicating that in the presence of DITPA,

IUGR is playing a role in elevating astrocyte numbers in the cerebellar lobules. Astrocytes treated

with TH (T3) secrete growth factors which amplify neuronal proliferation (Martinez and Gomes,

2005), therefore it would be of interest to investigate whether treatment with TH analogue DITPA in

combination with the effects of IUGR is causing a similar response. Future studies should investigate

the levels of growth factors in the cerebellum following DITPA administration, as well as

proliferation of neuronal populations, such as granule cells in the EGL.

4.5.5 DITPA increased Purkinje cell linear density in early developing cerebellar

lobules.

Purkinje cells are the primary output neuron of the cerebellum and aid in the coordination of sensory

input and motor control. In the present study Purkinje cell somal area, linear density, or areal density

was not affected in IUGR rats at P14. This was an unexpected result, as Purkinje cell numbers are

reduced in the cerebellum of foetal IUGR guinea pigs and this result persists to 1 week of age (Mallard

et al., 2000). The conflicting results between studies are likely due to the different gestation lengths

of species used, and therefore the different duration of placental insuffuceny induced via vessel

ligation. In the present study DITPA treatment significantly increased the linear density of Purkinje

cells (cells/mm) in the early developing lobules, irrespective of group (i.e. in both control and IUGR

pups). Purkinje cells undergo a period of natural cell death during normal cerebellar development

(Fan et al., 2001), an important process that regulates the final cell number (Baader et al., 1996, Vogel

and Herrup, 1993, Wetts and Herrup, 1982, Zanjani et al., 1996). Although not known in the rat, in

the mouse cerebellum, which has a similar growth trajectory, this occurs between P1 to P5 (Ghoumari

et al., 2000). Therefore it is possible that DITPA prevents natural cell death of Purkinje cells in early

developing cerebellar lobules, thus resulting in an increased density. It is unclear why this result was

only seen in the early developing lobules, or what the functional outcomes of this change would be.

Due to the design of this study, behavioural testing at P14 was not possible, however it would be

relevant for future studies to examine the effect of DITPA administration motor control and

coordination at a later time-point.


This study found that myelination, when assessed by analysing area of coverage of MBP-IR, was not

different in IUGR compared to control pups at P14. This was a surprising result, and it was thought

that a stress response might be at play. As discussed in Chapter 3 (Section 3.8.4) a limitation of this

study is the absence of absolute control groups, i.e. control and IUGR pups without intervention (daily

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handling, weighing, injecting) that would help elucidate the effects of IUGR vs. control without the

potential confounding variable of stress. Another limitation of this study is that only males were

assessed. This was so that the outcomes of the present study using longer-term DITPA treatment

could be compared to those of the IUGR study using short-term DITPA. However it should be

acknowledged that it is important to examine the effects of both IUGR and DITPA in both sexes; the

cerebellum of female rats from all experimental groups was also collected and these will be studied

at a later point in time.

4.5.7 Conclusion

The present study showed that DITPA administration in both IUGR and control pups did not affect

overall cerebellar morphology at P14. However DITPA administration decreased linear density of

BG fibres in the late developing cerebellar lobules of control pups, increased linear density of

Purkinje cells in early developing cerebellar lobules in both control and IUGR pups, and resulted in

microgliosis in the late developing lobules of IUGR pups. This study showed that the presence of

IUGR did not cause injury in the developing cerebellum, and importantly the rat cerebellar WM was

not affected at P14. This finding may serve for interspecies comparisons in order to elucidate

pathways of injury and resistance against IUGR. This study also showed that DITPA administration

from P1-P14 provided unfavourable effects in control animals, however it is useful to determine

whether DITPA administration causes harm to the developing cerebellum in general. The longer-term

neurological consequences of these effects are yet to be determined. Please see thesis Chapter 6

(Section 6.5) for future research directions, including investigating an alternate route of DITPA

administration and behavioural studies.

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5 Assessment of neonatal

growth and wellbeing measures

following DITPA therapy in

the IUGR rat.

5.1 Preamble

The overall aim of this thesis is to examine the potential therapeutic benefits of DITPA treatment on

brain development in IUGR. As described in Chapter 3, DITPA treatment in IUGR pups improved

cerebral myelination at P14, evident by elevated MBP-IR staining in the cortical layer VI when

compared to IUGR pups treated with saline. In IUGR pups DITPA did not cause inflammation or

injury, but did reduce area coverage of PLP-IR in some of the regions examined and reduced MBP-

IR of cortical processes, perhaps due to altered transport of MBP into the processes. In the cerebellum

(Chapter 4), DITPA administration in IUGR pups did not affect overall cerebellar development at

P14, but did increase linear density of Purkinje cells in the early developing cerebellar lobules. An

increase in the density of microglia was found in the late developing lobules in IUGR + DITPA pups

compared to IUGR + saline. Having established some of the potential impacts of longer-term DITPA

on the IUGR brain, it is essential to determine if DITPA has any negative off-target effects on the

postnatal development of these pups. In this Chapter, body weight, brain weight, body composition

(via DEXA), thyroid function (plasma FT3 and FT4), and plasma liver enzymes (ALT and ALP) as

well as cholesterol were assessed in both males and females at P14.

5.2 Introduction

The IUGR foetus possesses an adaptive mechanism, known as the ‘foetal brain-sparing effect’,

whereby in response to placental insufficiency, there is vasodilatation of the foetal cerebral circulation

to ‘protect’ the brain. However this typically occurs at the expense of other organs like the kidneys

and liver, which remain growth restricted (Miller et al., 2016). Despite this ‘brain-sparing’

phenomenon, IUGR babies still display significant neurodevelopmental sequelae including

behavioural deficits and cognitive impairment, and a relationship has been found between reduced

regional and total brain volumes of IUGR foetuses, neonates, children and adolescents (Padilla et al.,

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2014b, Tolsa et al., 2004, Dubois et al., 2008). A number of mechanisms have been attributed to the

brain injury that occurs in IUGR. Of relevance to this thesis, deficits in the cerebral TH transporter

MCT8 have been found in the brain of IUGR neonates compared to their normally grown counterparts

(Chan et al., 2014). TH is an essential regulator of neuronal (Thompson and Potter, 2000) and OL

development in the brain (Lee et al., 2017, Rodriguez-Pena, 1999). In the IUGR brain, reduced

expression of MCT8 transporters likely results in decreased TH signalling to cells, which may lead

to the impaired OL development and reduced myelination as seen clinically (Chan et al., 2014) (see

Chapter 1, Section 1.8 for role of TH in brain development). TH circulates in the blood as T4 and

T3, and an active fraction of these are called ‘free’ (FT3, FT4) and can be measured via blood plasma

assays (see Chapter 1, Section 1.7.1 for TH signalling). FT3 is 5 times more biologically active than

T4, and once inside the cell T4 undergoes a process called deiodination, whereby enzymes known as

iodothyronine deiodinases (Dio1, Dio2, Dio3) (Gereben et al., 2008) remove iodine atoms from T4

(Visser et al., 1988), producing an active T3 state. T3 can then enter the nucleus and bind to nuclear

TH receptors and regulate the expression of specific genes in that cell.

TH signalling in the body relies on a negative feedback mechanism to maintain homeostasis. For

example, if circulating TH levels in the blood become too low, the hypothalamus secretes TRH which

stimulates the pituitary gland to secrete TSH, and this signals the thyroid gland to release more T3

and T4 into the blood stream, thus normalising circulating TH levels. Conversely, if circulating TH

levels are too high, the hypothalamus down regulates TRH secretion, ultimately leading to the thyroid

gland releasing less TH into circulation. If homeostasis is disturbed, this feedback loop compensates

for the disruption (Dietrich et al., 2012). As the TH transporter MCT8 is deficient in the IUGR brain

(Chan et al., 2014), conventional TH therapy will likely be ineffective, and therefore new therapies

must be explored. One such therapy is the TH analogue DITPA, which does not require MCT8 to

enter cells in the brain (Ferrara et al., 2015).

While the clinical safety profile of DITPA has been established in children with MCT8 mutation

(Verge et al., 2012), adults with cardiovascular disease (Goldman et al., 2009, Ladenson et al., 2010),

and in MCT8 knockout mice (Ferrara et al., 2015), it has not yet been established in IUGR neonates.

A previous study by our group examined DITPA treatment from P1 to P6 as a reparative treatment

following IUGR for cerebral hypomyelination (Azhan, A., unpublished thesis, 2019). This study also

assessed off-target effects of DITPA, examining body, brain, liver and kidney weight, as well as body

composition at P7, and found that DITPA did not affect these measures. Reduced levels of MCT8

mRNA were also found in IUGR pups compared to controls at P7, however there were no differences

at P14 (Azhan, A., unpublished thesis, 2019). While DITPA did not result in off-target effects

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following short-term treatment (6 days), the impact of longer-term DITPA treatment still needs to be

examined. There is currently no clinical treatment for IUGR, with the only intervention being to

deliver the baby preterm to remove it from the intrauterine environment and the insufficient placenta;

this could occur any time between 24 and 40 weeks GA. Once removed from the unfavourable

intrauterine environment, DITPA therapy would ideally be given to the baby from the time of delivery

until MCT8 levels had normalised. This information is currently unavailable for the IUGR infant,

however in the IUGR rat, MCT8 mRNA levels normalised to control levels by P14 (Azhan, A.,

unpublished thesis, 2019) an age equivalent to brain development in a human baby at term age (~40

weeks GA) (Semple et al., 2013).

This Chapter aimed to investigate the effects of daily DITPA administration from P1 to P13 on

neonatal health and wellbeing, including body and organ weights, body morphometry, body

composition, thyroid function, liver function, and cholesterol levels in IUGR pups at P14. The

hypothesis was that DITPA administration in IUGR rat pups from P1 to P13 would not have a

negative impact on any of these measures of neonatal health and wellbeing.

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5.3 Methodology

For comprehensive methodology regarding animal welfare, species, surgical procedure, drug

treatment, post-mortem and tissue collection, please refer to Chapter 2 (Sections 2.2 to 2.8).

5.3.1 Overview of animal work

At day 18 of pregnancy (term = 22 days), rats underwent BUVL (n = 31 dams) or sham surgery (n =

16 dams) to generate IUGR or control pups. DITPA (0.5mg/100g; i.p.) or saline (equivalent volume)

was administered daily from P1 to P13 to IUGR (IUGR + DITPA: n = 14 dams, 67 pups; IUGR +

saline: n = 17 dams, 60 pups) and control (control + DITPA, n = 8 dams, 49 pups; control + saline: n

= 8 dams, 47 pups) pups. Not all BUVL/sham surgeries were successful (i.e. they did not yield IUGR

or control pups) and these were not included in the study (Figure 5.1). Body weight was measured

daily from P1 to P14, and brain weight, body composition (via DEXA), thyroid function (plasma FT3

and FT4), plasma liver enzymes (ALT and ALP) and plasma cholesterol were assessed at P14.

5.3.2 Body and organ weights

Pups were weighed and sexed on P1 to avoid the dam becoming stressed and rejecting pups

immediately after birth (see Chapter 2, Section 2.6.1). Pups were weighed daily (prior to saline or

DITPA treatment) from P1 to P13, and on P14 prior to euthanasia. At P14, pups’ head and hip

circumference as well as crown-to-rump length (dorsal distance from crown of skull to base of spine)

Figure 5. 1 Overview of animals used in Chapter 5. A total of 223 P14 rat pups were used in this study.

This included both male and female control and IUGR pups, treated with DITPA or saline. The number

of dams is shown for each surgery group (sham/BUVL). Not all BUVL/sham surgeries were successful; the

number of successful litters used in experimentation is shown. n = number of animals.

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were measured using a measuring tape (mm). All measurements were conducted following euthanasia

to minimise measuring variability caused by movement. Kidney, liver and brain weight (whole, and

dissected into hemispheres, cerebellum, pons, medulla and cervical spinal cord) were recorded for

each pup at post-mortem on P14. Livers from pups that were not perfused were collected and then

frozen in liquid nitrogen and stored at -80°C for later assessment of Dio1 mRNA (see Section 5.3.5).

5.3.3 Analysis of body composition using dual-energy x-ray absorptiometry

(DEXA)

At P14, carcasses (excluding brain, kidney and liver) of pups were frozen at -4°C and transported on

dry ice to the Monash University DEXA body composition and bone density analysis suite (Clayton,

Victoria, Australia). DEXA uses low-energy x-rays from two different sources (improves reliability)

to measure the density and composition of bone, fat and soft tissue. Pups were placed in the supine

position on the ‘tray’ of the DEXA machine and scanned to determine: total bone area, bone mineral

density and composition, lean tissue mass, fat mass and percentage of total body fat. The head of the

animal was excluded from this analysis (Figure 5.2). All readings were recorded as grams per cm2

except for % body fat which was recorded as a proportion (%).

Figure 5. 2 Image of P14 rat taken using dual-energy x-ray absorptiometry (DEXA). Following

euthanasia, bodies of pups were scanned using a DEXA machine. Body composition measures such as total

bone area, bone mineral density and composition, lean tissue mass, fat mass and percentage of total body fat

were analysed. Green circle = the skull, was not included in this analysis; blue trace = outline of the pup’s body;

yellow trace = bone (note: not all the bone is traced in yellow, however the DEXA machine does account for

all bone mass). Image not to scale.

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5.3.4 Analysis of blood plasma

At P14, blood was collected from the right ventricle of each pup immediately following euthanasia

using a 26-gauge needle and 1mL syringe, placed into Eppendorf tubes containing heparin (2μL, 5000

IU in 5mL; Pfizer, Australia) to prevent clotting, and kept on ice until they were centrifuged at 22°C

and 2600rpm for 10 minutes (Eppendorf centrifuge 5819 R, Hamburg, Germany) to separate the

blood plasma from the red blood cells and other constituents. Following centrifuging, the plasma

layer was collected using a 1ml syringe (minimum volume required = 300μL), and immediately

frozen at -80°C. These plasma samples were later used in hospital standard plasma assays (Monash

Pathology, Clayton, Victoria, Australia) to assess thyroid function (free T3 and T4), liver enzymes

(ALT and ALP) and cholesterol (protocols in Appendix 6).

5.3.5 Analysis of Dio1 in the Liver

RNA extractions

Ribonucleic Acid (RNA) was extracted from liver samples (control + saline: n= 6; control + DITPA:

n= 6; IUGR + saline: n= 7; IUGR + DITPA: n = 7). Frozen liver samples were dry crushed into a fine

powder using a pestle and mortar, pre-cooled with dry ice and liquid nitrogen. Approximately 20mg

of liver was weighed into pre-chilled 2ml Eppendorf tubes with a stainless-steel bead. RNA was

extracted at room temperature using the PureLink RNA Mini Kit (Invitrogen, MA, USA) as per

manufacturer’s instructions. Briefly, 600μL lysis buffer containing 0.01% β-mercaptoethanol was

added to each sample before homogenising at 50Hz for 2 minutes using the TissueLyser LT (Qiagen,

Hilden, DE). Samples were centrifuged at 12,000g to remove debris and supernatant was transferred

to a fresh tube. One part ethanol was added to each sample and vortexed to combine. Samples were

added to the spin columns provided in the kit and centrifuged at 12,000g for 20 seconds to elute buffer

and bind RNA to the column. The flow-through was discarded, with the process repeated until all

sample was bound. Following this, liver samples were washed with 700μL of Wash Buffer 1 and

centrifuged 12,000g for 20 seconds and flow-through discarded. Samples were then washed twice

with 500μL of Wash Buffer 2, centrifuged at 12,000g for 20 seconds and flow-through discarded;

samples were dried by centrifuging for 2 minutes at 12,000g. Spin columns were then transferred into

1.5mL collection tubes, with 100μL of MilliQ water (Merck Millipore, Darmstadt, DE) and incubated

for 1 minute at room temperature. Samples were then centrifuged at >12,000g to elute RNA. RNase

inhibitor was added to each sample the prevent degradation.

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RNA concentrations, purity and quality

RNA concentrations for liver samples were detected using the NanoDrop One spectrophotometer

(Thermo Fisher Scientific, Scoresby, Victoria, Australia). The fibre optic point was calibrated using

2µl of MillliQ water. 2 µl of each sample were then individually read and calculated by the machine.

Nucleic acid concentration (ng/µl), absorbance (260nm) and RNA purity (A260/A280 and A260/A230

ratios) were recorded. RNA was assessed to be of good quality if the RNA concentration was above

100 ng/µl, with the A260/A280 ratio 2, and A260/A230 ratio 2.2. Deviations from these expected

values indicate contamination by proteins, organic compounds or low RNA concentrations. Samples

that gave an indication of contamination via unexpected deviations in these measures were discarded

and the RNA extraction was repeated. Integrity of RNA was assessed via agarose gel electrophoresis.

Briefly, the apparatus was soaked in RNase Away (Invitrogen, MA, USA) for 30 minutes to remove

RNases. Agarose powder (0.3 g) was mixed with 30 ml 1x TAE (Tris-base, acetic acid, 0.5M EDTA

buffer), and the mixture was then heated for approximately 30 seconds (until clear) to make a 1%

agarose gel. SYBR safe (Invitrogen, MA, USA) was added to the gel at 1:10,000 for RNA detection.

The gel was poured into the mould and allowed to set for 30 to 60 minutes. Approximately 5μL of

liver sample was mixed with 1μL of 6x TrackIt Cyan/Orange Loading Buffer (Invitrogen, MA, USA),

and loaded into wells. TrackIt 1 Kb Plus DNA Ladder (Invitrogen, MA, USA) was also loaded to

track RNA banding. The gel underwent electrophoresis at 100 V for 60 minutes, or until the yellow

dye-front reached the end of the gel. The agarose gel was imaged at 530 nm (green light) using the

FluorChem Q (Alpha Innotec GmbH, Kasendorf, Germany). Quality RNA was determined by the

lack of sample smearing through the gel, and the presence of the 18S and 28S bands at a 1:2 ratio

signal intensity.

Reverse transcription of liver samples

Liver samples were reverse transcribed to complementary DNA (cDNA) using the High Capacity

cDNA Reverse Transcription Kit (Applied Biosystems, CA, USA) as per manufacturer’s instructions.

All reagents and samples were kept on ice. One tube contained the sample with reverse transcriptase

to make cDNA (labelled RT+) and one tube contained all of the same reagents, however included

MilliQ water instead of the reverse transcriptase enzyme. This tube (labelled RT-) acted as a control

for DNA contamination of the sample. Briefly, sample RNA and MilliQ was pipetted into RT+ and

RT- tubes to a total amount of 1μg RNA in 10μL. Reaction buffers were created containing 2μL 10X

RT Buffer, 0.8µl 25X dNTP mix, 2µl 10X Random Primers, 1 µl of MultiScribe Reverse

Transcriptase and 4.2µl MilliQ water per sample. The RT-mixture consisted of the same components

except MultiScribe Reverse Transcriptase which was replaced with MilliQ water. 10µl of the reaction

mixture was added to each Eppendorf tube. Samples were gently mixed then spun down before

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incubation in a heat-block (Ratek, Boronia, Victoria, Australia). Briefly, RNA strands were denatured

for 10 minutes at 25°C before cDNA synthesis of RNA strands at 37°C for 2 hours. cDNA synthesis

was terminated by incubation for 5 minutes at 85°C. Samples were quenched on ice, before being

stored at -20°C until use. The final concentration obtained for each sample was 50ng/μL.

Real-time PCR (qPCR) for Dio1 using Taqman probes

TaqMan (Applied Biosystems, CA, USA) was used to analyse PCR products for all genes examined.

All liver cDNA samples were diluted to a working concentration of 10ng/μL. Relative expression of

Dio1 was determined via the 2-ΔΔCT method with β2 microglobulin (B2M) used as the housekeeping

gene (See table 5.1 for Assay ID).

Table 5. 1 Gene assay ID’s.

Master mixes containing TaqMan Fast Advanced Master Mix (Applied Biosystems, CA, USA), 1x

TaqMan Gene Expression Assay probes (Dio1 or B2M) and MilliQ water were made for all samples.

All RT+ samples were run in duplicate, with its corresponding RT- sample also run to confirm the

removal of DNA contamination; no-template controls were also run in triplicate on the plate to assess

for contamination of the PCR plate or master mixes. Samples were pipetted into a MicroAmp Optical

384-well plate (Applied Biosystems, CA, USA) at 1μL (10ng/μL), and 9μL of TaqMan master mix

was added to each well. The plate was sealed with MicroAmp Optical Adhesive Film (Applied

Biosystems, CA, USA), gently mixed and centrifuged for 20 seconds at 200g in a Heraeus refrigerated

bench-top multifuge (Heraeus 3XR multifuge, Thermo Fisher Scientific, MA, USA) to combine all

reagents in the bottom of the wells and to remove air bubbles that could affect the reading. Real time

PCR was performed on the QuantStudio Flex Real-Time PCR Systems (Applied Biosystems, CA,

USA) and analysed using the QuantStudio Real-Time PCR Software v1.6.1 (Applied Biosystems,

CA, USA). Plates were placed in the real-time machine and selected for detection of gene of interest.

Cycle threshold (Ct) for detection was manually set for exponential phase, at 1.281 and 0.508

fluorescence for Dio1 and B2M, respectively.

2-ΔΔCT analysis of Dio1 relative expression

The Ct value for the duplicates of each liver sample was averaged. The ΔCt was calculated by

subtracting the B2M average Ct from the Dio1 average Ct. To calculate the ΔΔCt value, the ΔCt of

Gene Species Assay ID Dye

Dio1 Rat Rn00572183_m1 FAM-MGB

B2M Rat Rn07310889_g1 VIC-MGB

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the control sample group (control + saline) was subtracted from the sample ΔCt of each sample.

Finally, the relative fold change difference of samples was collected by taking the –ΔΔCt value to the

power of 2 (2-ΔΔCt).

ΔCtsample = CtDio1 – CtB2M

ΔΔCt = ΔCtsample – ΔCtcontrol group

Relative fold change = 2-ΔΔCt

5.3.6 Statistical analysis

All statistical analyses on data in this Chapter were performed using IBM SPSS® (version 25; SPSS

Inc.; IBM Corporation, Armonk, NY, USA), and graphs were created using Graphpad Prism

statistical software (version 8.0, Graphpad Software Inc., La Jolla, CA, USA). Prior to analyses,

outliers were removed using the Grubb’s test to determine a significant outlier (significance = p

<0.05). The data from all pups in every litter were included in this study (See Figure 5.1). It was

necessary to account for variation between litters, when determining the impact of IUGR and DITPA

treatment. A linear mixed model was used to examine the association between treatment (saline or

DITPA) and group (control or IUGR) outcomes, and whether this differed with sex. The main effects

plus three interaction terms (group*sex, group*treatment, group*treatment*sex) were included in the

model as fixed effects, and a random intercept for mother was included to control for correlations

between pups within a litter. Statistical significance was determined using the type III test of fixed

effects for interactions (*). Data was considered significant at p < 0.05. A Bonferroni correction was

used for multiple post-hoc comparisons within group, treatment and sex. All main effects data are

presented as the ratio of residual variances (F) and statistical significance (p-value). All post-hoc data

are presented as [mean comparative difference (95% confidence interval); p-value], and in graphs the

data is shown as Mean SEM except for body measurements and organ weight data which are

presented as Mean standard deviation (SD), as required for IUGR classification.

Supplementary data tabulated in Appendix 5 is shown as Mean SEM or SD for weights and

measurements.

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5.4 Results


Body weights at P1, P7 and P14

Linear mixed model analysis showed no interactions between group, treatment or sex on body weight

(g) of pups at P1, P7 or P14. There was a main effect of group and treatment on body weight of pups

at P1 (treatment: F1/206 = 4.91, p < 0.0001; group: F1/206 = 406.5, p < 0.0001), an effect of only group

on body weight of pups at P7 (F1/206 = 244.8, p < 0.001), and an effect of both group and treatment at

P14 (treatment: F1/200 = 6.43, p = 0.01; group: F1/200 = 126.3, p < 0.0001).

Post-hoc analysis showed that body weight was reduced in IUGR + saline pups compared to control

+ saline pups at P1 [1.69 (1.43, 1.94) p < 0.0001; Figure 5.3 A], P7 [5.05 (4.21, 5.89) p < 0.0001;

Figure 5.3 D] and P14 (7.03, 5.53 – 8.53, p < 0.0001; Figure 5.3 G). This was also true when males

and females were analysed separately at P1 [male: 1.85 (1.50, 2.20); p < 0.0001; female: 1.53 (1.17,

1.88) ; p < 0.0001; Figure 5.3 B, C], P7 [male: 5.48 (4.30, 6.66); p < 0.0001; female: 4.61 (3.42,

5.80); p < 0.0001; Figure 5.3 E, F] and P14 [male: 7.65 (5.56, 9.75); p < 0.0001; female: 6.37 (4.22,

8.52); p < 0.0001; Figure 5.3 H, I].

Post-hoc analysis also showed increased body weight in control + DITPA pups compared to control

+ saline at P1 in sexes combined [0.33 (0.63, 0.60); p = 0.02; Figure 5.3 A], and in females only when

sexes were analysed separately at P1 [0.57 (0.17, 0.97); p = 0.005; Figure 5.3 C]. Body weight was

reduced in control + DITPA pups compared to control + saline pups in sexes combined at P14 [-2.28

(3.89, -6.61); p = 0.006; Figure 5.3 G], and in males only when sexes were analysed separately at P14

[2.75 (0.55, 4.97); p = 0.02; Figure 5.3 H]. Body weight was reduced in IUGR + DITPA pups

compared to control + DITPA pups at P1, P7 and P14 in sexes combined [P1: -1.95 (-2.21, -1.70); p

< 0.0001; P7: -1.80 (-2.15, -1.46); p < 0.0001; P14: -2.10 (-2.48, -1.73); p < 0.0001], and when sexes

were analysed separately; this was true in both males [P1: -1.80 (-2.15, -1.46); p < 0.0001; P7: -4.09

(-5.25, -2.93), p < 0.0001; P14: -4.79 (-6.88, -2.70); p < 0.0001; Figure 5.3 B,E,H] and females [P1:

-2.10 (-2.48, -1.73); p < 0.0001; P7: -4.87 (-6.13, -3.60); p < 0.0001; P14: -5.64 (-7.89, -3.39); p <

0.0001; Figure 5.3 C,F,I]. There was no difference in body weight between IUGR + DITPA and

IUGR + saline pups at P1, P7 or P14 in sexes combined, or when males and females were analysed

separately (p > 0.05; Figure 5.3 A-I).

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See Appendix 5 for Mean ± SD of all body weight data.

Liver and kidney weights at P14

Linear mixed modelling showed no significant interactions between group, treatment, or sex on liver

and kidney (left and right) weight (g) at P14 (Type III, p > 0.053). There was a main effect of group

and treatment on liver weight (treatment: F1/186 = 7.29; group: F1/186 = 64.5, p < 0.0001 both), as well

as the weight of the left kidney (treatment: F1/190 = 5.59, p = 0.02; group: F1/190 = 78.58, p < 0.0001),

and right kidney (treatment: F1/186 = 14.32; group: F1/186 =72.30; p < 0.0001 both), and a main effect

of sex on left kidney weight only (F1/190 = 5.58, p = 0.02) at P14.

Post-hoc analysis showed that liver weight was decreased in IUGR + saline compared to control +

saline pups in sexes combined [0.27 (0.19, 0.35); p < 0.0001; Figure 5.4 A], and this was true for both

Figure 5.3 Body weights (g) at postnatal day 1 (A-C), P7 (D-F), and P14 (G-I) in control and IUGR

pups treated with DITPA or saline. Data analysed using a linear mixed model with Bonferroni correction.

Values are presented as Mean ± SD. g = grams. * p < 0.05, ** p < 0.01, ****p < 0.0001. Pup numbers: control

+ saline: n = 34 male, n = 33 female; control + DITPA: n = 33 male, n = 27 female; IUGR + saline: n = 31

male, n = 18 female; IUGR + DITPA: n = 23 male, n = 24 female.

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males and females [males: 0.023 (0.12, 0.35); p < 0.0001; females: 0.31 (0.19, 0.43); p < 0.0001;

Figure 5.4 B, C]. Liver weight was reduced in control + DITPA female pups compared to control +

saline when sexes were analysed separately [0.16 (0.03, 0.30) p = 0.02; Figure 5.4 C] but there was

no difference in males, or when sexes were combined (Figure 5.4 A, B).

Post-hoc multiple comparisons also showed that kidney weight (left and right) was reduced in IUGR

+ saline pups when compared to control + saline pups in sexes combined [left kidney: 0.05 (0.05,

0.06); p < 0.0001; right kidney: 0.04 (0.03, 0.06); p < 0.0001; Figure 5.4 D, G], and this was true for

both males and females [male left kidney: 0.04 (0.02, 0.05); p < 0.0001; female left kidney: 0.05

(0.03, 0.07); p < 0.0001; male right kidney: 0.04 (0.02, 0.05); p < 0.0001; female right kidney: 0.05

(0.03, 0.07); p < 0.0001; Figure 5.4 E, F, H, I].

Left kidney weight was increased in IUGR + DITPA compared to IUGR + saline pups, when sexes

were combined [0.02 (0.002, 0.029); p = 0.023; Figure 5.4 D]. Right kidney weight was increased in

IUGR + DITPA compared to IUGR + saline pups when sexes were combined [0.022 (0.01, 0.03); p

< 0.0001; Figure 5.4 G], and in males and females when assessed separately [male: 0.025 (0.009,

0.042); p = 0.003; female: 0.019 (0.002, 0.03); p = 0.03; Figure 5.4 H, I]. Left and right kidney weight

was reduced in IUGR + DITPA pups compared to control + DITPA pups in sexes combined [left

kidney: -0.042 (-0.06, -0.021); p < 0.0001; right kidney: -0.034 (-0.05, -0.03); p < 0.0001; Figure 5.4

D, G], and this was true for both males and females [male left kidney: -0.045 (-0.06, - 0.03); p <

0.0001; female left kidney: -0.04 (-0.06, - 0.02); p < 0.0001; male right kidney: -0.03 (-0.05, -0.02);

p = 0.001; female right kidney: -0.038 (-0.06, - 0.02); p < 0.0001; Figure 5.4 E,F,H,I].

See Appendix 5 for Mean ± SD of liver and kidney weight data.

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5.4.2 Brain weights

Linear mixed modelling showed no interactions between group, treatment or sex on total brain weight

or weight of the cerebral hemispheres, cerebellum, pons, medulla (all in grams) or brain-to-body

weight ratio at P14. There was a main effect of group on total brain weight (F1/131 = 71.23, p < 0.0001)

and weight of the cerebral hemispheres (F1/87 = 30.62, p < 0.0001), cerebellum (F1/168 = 4.23, p =0.04),

pons (F1/164 = 9.67, p = 0.002), medulla (F1/164 = 9.32, p = 0.003) and brain-to-body weight ratio (F1/99

= 42.97, p < 0.0001). There was a main effect of treatment on pons weight (F1/164 = 5.07, p = 0.03)

and brain-to-body weight ratio (F1/190 = 4.50, p = 0.04), and a main effect of sex on total brain weight

only (F1/189 = 13.95, p < 0.0001).

Figure 5.4 Liver (A-C) and kidney (D-I) weights (g) at P14 in male and female control and IUGR pups

treated with DITPA or saline. Data analysed by linear mixed modelling with Bonferroni correction. Values

presented as Mean ± SD. g = grams, * p < 0.05, ** p < 0.01, *** p < 0.001, **** p < 0.0001. Pup numbers:

control + saline: n = 34 male, n = 33 female; control + DITPA: n = 33 male, n = 27 female; IUGR + saline: n

= 31 male, n = 18 female; IUGR + DITPA: n = 23 male, n = 24 female.

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Total brain weight at P14

Post-hoc analysis showed that at P14, total brain weight (g) was reduced in IUGR + saline compared

to control + saline pups in sexes combined [0.93 (0.062, 0.124); p < 0.0001; Figure 5.5 A], and this

was true for both males and females when analysed separately [males: 0.01 (0.05, 0.14); p < 0.0001;

females: 0.09 (0.05, 0.13); p < 0.0001; Figure 5.5 B, C]. Total brain weight was reduced in IUGR +

DITPA pups compared to control + DITPA pups in sexes combined [-0.011 (-0.014, -0.07); p <

0.0001; Figure A], and when analysed separately, this was seen in both males and females [males: -

0.12 (-0.16, -0.07); p < 0.0001; females: -0.09 (-0.01, -0.05); p < 0.0001; Figure 5.5 B,C]. DITPA

treatment did not affect total brain weight in IUGR or control pups compared to saline (Figure 5.5 A-

C).

Brain-to-body weight ratio at P14

Post-hoc analysis showed that IUGR + saline pups had an increased brain-to-body weight ratio (g/g)

compared to control + saline pups at P14 [0.006 (0.004, 0.009); p < 0.0001; Figure 5.5 D], and this

was true for both males and females [males: 0.007 (0.004, 0.10); p < 0.0001; females: 0.006 (0.003,

0.008); p < 0.0001; Figure 5.5 E, F]. Brain-to-body weight ratio was increased in control + DITPA

compared to control + saline pups in sexes combined [0.003 (0.001, 0.005); p = 0.01; Figure 5.5 D],

and when analysed separately this was seen in males [0.004 (0.001, 0.007); p = 0.01; Figure 5.5E]

but not females. Brain-to-body weight ratio was increased in IUGR + DITPA pups compared to

control + DITPA pups when sexes were combined [0.0004 (0.002, 0.006); p < 0.0001; Figure 5.5 D],

and in males and females when assessed separately [males: 0.0003 (0.0, 0.006); p = 0.03; females:

0.005 (0.002, 0.008); p = 0.001; Figure 5.5 D, F]. DITPA did not affect brain-to-body weight ratio in

IUGR pups compared to saline administration (Figure 5.5 D-F).

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Cerebral hemisphere weight at P14

Post-hoc analysis showed that cerebral hemisphere weight was reduced IUGR + saline compared to

control + saline pups when sexes were combined [-0.09 (-0.14, -0.45); p < 0.0001; Figure 5.6 A] and

when sexes were analysed separately [males: -0.10 (-0.03, -0.16); p = 0.003; females: -0.09 (-0.02, -

0.15); p – 0.007; Figure 5.6 B, C]. Hemisphere weight was reduced in IUGR + DITPA pups compared

to control + DITPA pups when sexes were combined [-0.011 (-0.15, - 0.06); p < 0.0001; Figure 5.6

A], and in males and females separately [males: -0.11 (-0.17, -0.04); p = 0.002; females: -0.11 (-0.17,

- 0.04); p = 0.002; Figure 5.6 B, C]. There was no difference in cerebral hemisphere weight in IUGR

or control pups treated with DITPA compared to saline (Figure 5.6 A); this was also true when males

and females were assessed separately (Figure 5.6 B,C).

Cerebellum weight at P14

Post-hoc analysis revealed that there was no difference in cerebellar weight (g) between IUGR +

saline pups and control + saline pups in sexes combined (Figure 5.6 D), or in males and females when

analysed separately (Figure 5.6 E, F). Cerebellar weight was decreased in IUGR + DITPA pups

compared to control + DITPA pups in sexes combined [-0.02 (-0.03, -0.004); p = 0.012; Figure 5.6

D], but not in males and females when analysed separately (Figure 5.6 E, F). There was no difference

Figure 5.5 Total brain weight (g) (A-C) and brain-to-body weight ratio (D-F) at P14 in male and female

control and IUGR pups treated with DITPA or saline. Data analysed by linear mixed modelling and

Bonferroni correction. Values presented as Mean ± SD. g = grams, * p < 0.05, *** p < 0.001, **** p < 0.0001.

Pup numbers: control + saline: n = 34 male, n = 33 female; control + DITPA: n = 33 male, n = 27 female;

IUGR + saline: n = 31 male, n = 18 female; IUGR + DITPA: n = 23 male, n = 24 female.

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in cerebellar weight in IUGR or control pups treated with DITPA or saline, in sexes combined (Figure

5.6 D) or when males and females were assessed separately (Figure 5.6 E, F).

Pons weight at P14

Post-hoc multiple comparisons showed that pons weight (g) was reduced IUGR + saline pups

compared to control + saline pups when sexes were combined [-0.009 (- 0.14, - 0.004); p = 0.001;

Figure 5.6 G] and assessed separately [male: -0.010 (-0.017, -0.002); p = 0.01; female: -0.008 (0.001,

0.015); p = 0.03; Figure 5.6 H, I]. Pons weight was reduced in control + DITPA compared to control

+ saline pups in sexes combined [-0.007 (-0.013, 0.002); p = 0.008; Figure 5.6 G], and in males [0.010

(0.002, 0.01); p = 0.01; Figure 5.6 H] but not females (Figure 5.6 I) when analysed separately.

Medulla weight at P14

Post-hoc multiple comparisons showed that there was no difference in the weight (g) of the medulla

in control + saline compared to IUGR + saline pups at P14 when sexes were combined (Figure 5.6 J)

or assessed separately (Figure 5.6 K, L). The weight of the medulla weight was reduced in IUGR +

DITPA compared to control + DITPA pups when sexes were combined [- 0.006 (-0.002, - 0.011); p

= 0.006; Figure 5.6 J], and in males [-0.008 (- 0.001, -0.014); p = 0.02; Figure 5.6 K], but not females

(Figure 5.6 L) when sexes were analysed separately. There was no difference in medullar weight in

IUGR or control pups treated with DITPA compared to those treated with saline (Figure 5.6 J, K, L).

See Appendix 5 for Mean ± SD of all brain weight data.

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5.4.3 Body morphometry

Linear mixed modelling analysis showed no interactions between group, treatment or sex on crown-

to-rump length (CRL), head circumference or hip circumference (Type III, p > 0.05). There was a

main effect of group on CRL (F1/199 = 17.56, p < 0.0001), head circumference (F1/199 = 4.82, p = 0.03)

and hip circumference (F1/198 = 12.91, p < 0.0001), but no main effects of treatment or sex.

Figure 5.6: Weight (g) of the cerebral hemispheres (A-C), cerebellum (D-F), pons (G-I) and medulla (J-

L) in male and female control and IUGR pups at P14 treated with DITPA or saline. Data analysed by

linear mixed modelling and Bonferroni correction. Values presented as Mean ± SD. g = grams, * p < 0.05, **

p < 0.01, *** p < 0.001, **** p < 0.0001. Pup numbers: control + saline: n = 34 male, n = 33 female; control

+ DITPA: n = 33 male, n = 27 female; IUGR + saline: n = 31 male, n = 18 female; IUGR + DITPA: n = 23

male, n = 24 female.

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Crown-to-rump-length (CRL) at P14

Post-hoc multiple comparisons showed that at P14 CRL (mm) was decreased in IUGR + saline

compared to control + saline pups when sexes were combined [-7.38 (-11.31, 3.45); p < 0.0001;

Figure 5.7 A] and analysed separately [males: -9.45 (-14.82, - 4.09); p = 0.01; females: -5.30 (-10.58,

-0.021); p = 0.049; Figure 5.7 B, C]. CRL was decreased in IUGR + DITPA pups compared to control

+ DITPA pups in sexes combined [-5.24 (-9.13,-1.35); p= 0.009; Figure 5.7 A], and males only [-

5.29 (-10.48, -0.10); p = 0.046; Figure 5.7 B]. There was no difference in CRL between IUGR or

control pups treated with DITPA compared to those treated with saline (Figure 5.7 A, B, C).

Head circumference at P14

Post-hoc analysis showed that head circumference (mm) was reduced in IUGR + DITPA pups

compared to control + DITPA pups when sexes were combined [-3.44 (-6.02, - 0.87); p = 0.009;

Figure 5.7 D], and in males [-4.13 (-7.60, - 0.66); p = 0.02] but not females (Figure 5.7 F). There was

no difference in head circumference between control + saline and IUGR + saline pups, and this was

true for both males and females (Figure 5.7 D-F). There was no difference in head circumference in

IUGR or control pups treated with DITPA compared to those treated with saline, when sexes were

combined or assessed separately (Figure 5.7 D-F).

Hip circumference at P14

Post-hoc analysis showed that hip circumference (mm) was reduced in IUGR + saline pups compared

to control + saline pups when sexes were combined [- 4.49 (- 7.61, - 1.37); p = 0.005; Figure 5.7 G]

and in males [-7.62 (-11.92, - 3.31); p = 0.001; Figure 5.7 H] but not females (Figure 5.7 I). Hip

circumference was reduced in IUGR + DITPA pups compared to control + DITPA pups in sexes

combined [-3.59 (-6.75, - 0.44); p = 0.026; Figure 5.7 G]. There was no difference in hip

circumference in IUGR or control pups treated with DITPA compared to those treated with saline

(Figure 5.7 G, H, I).

See Appendix 5 for Mean ± SD of body morphometry data.

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5.4.4 Body composition (DEXA)

Linear mixed modelling analysis showed no interactions between group, treatment or sex on bone

mineral density (g/cm2), bone mineral content (g), total bone area (cm2), lean tissue mass (g), fat mass

(g) or percentage of total body fat (%)(Type III, p > 0.05). There was an overall effect of group on

bone mineral density (F1/149 = 27.9), mineral content (F1/82 = 42.9), total bone area (F1/156 = 48.5), lean

tissue mass (F1/156 = 62.8), fat mass (F1/156 = 48.5) (p < 0.0001 for all), but not % body fat. There was

a main effect of treatment on bone mineral density only (F1/149 = 4.1, p = 0.006).

Figure 5.7: Crown-to-rump length (A-C), head circumference (D-F), and hip circumference (G-I) (mm)

in male and female control and IUGR pups at P14 treated with DITPA or saline. Data analysed by linear

mixed modelling and Bonferroni correction. Values presented as Mean ± SD. mm = millimetres, * p < 0.05,

** p < 0.01, *** p < 0.001, **** p < 0.0001. Pup numbers: control + saline: n = 34 male, n = 33 female;

control + DITPA: n = 33 male, n = 27 female; IUGR + saline: n = 31 male, n = 18 female; IUGR + DITPA: n

= 23 male, n = 24 female.

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Bone mineral density at P14

Post-hoc multiple comparisons showed that bone mineral density (g/cm2) was reduced in IUGR +

saline compared to control + saline pups in sexes combined [-0.003 (-0.06, -0.01); p = 0.005; Figure

5.8 A], and when analysed separately, in males [-0.003 (-0.007, 0); p = 0.002; Figure 5.8 B] but not

females (Figure 5.8 C). Bone mineral density was increased in control + DITPA pups compared to

control + saline pups in sexes combined [-0.003 (-0.005, -0.00009); p = 0.01; Figure 5.8 A] and in

females [0.004 (0.00014, 0.008); p = 0.05; Figure 5.8 C], but not males (Figure 5.8 B). There was no

difference in bone mineral density between IUGR + DITPA pups compared to IUGR + saline pups

(Figure 5.8 A, B, C). Bone mineral density was increased in control + DITPA compared to IUGR +

DITPA pups in sexes combined [0.005 (0.003, 0.0008); p < 0.0001; Figure 5.8 A] and in males and

females when analysed separately [males: 0.004 (0.001, 0.007); p = 0.03; females: 0.07 (0.004,

0.011); p < 0.0001; Figure 5.8 B, C].

Bone mineral content at P14

Post-hoc multiple comparisons revealed that bone mineral content (g) was reduced in control + saline

vs IUGR + saline pups when sexes were combined [-0.10 (-0.15, - 0.06); p < 0.0001; Figure 5.8 D],

and in males and females when analysed separately [males: -0.9 (-0.15, -0.03); p = 0.002; females: -

0.11 (-0.17, -0.05); p = 0.001; Figure 5.8 E, F]. Bone mineral content was decreased in IUGR +

DITPA pups compared to control + DITPA pups in sexes combined [-0.10 (-0.15, -0.06); p < 0.0001;

Figure 5.8 D] and in males and females when analysed separately [-0.08 (-0.14, -0.03); p = 0.005;

females: -0.12 (-0.19, - 0.06); p < 0.0001; Figure 5.8 E, F]. There was no difference in bone mineral

content in IUGR or control pups treated with DITPA compared to those treated with saline (Figure

5.8 D, E, F).

Total bone area at P14

Post-hoc analysis showed that total bone area (cm2) was reduced in IUGR + saline compared to

control + saline pups when sexes were combined [-2.26 (-3.16, -1.36); p < 0.0001; Figure 5.8 G], and

in males and females when analysed separately [males: -2.05 (-3.27, -0.82); p = 0.001; females: -2.47

(-3.79, -1.16); p < 0.0001; Figure 5.8 H, I]. Total bone area was reduced in IUGR + DITPA compared

to control + DITPA pups in sexes combined [-2.26 (-3.17, -1.35); p < 0.0001; Figure 5.8 G], and

males and females when analysed separately [males: -1.69 (-2.88, -0.51); p = 0.005; females: -2.83 (-

4.21, -1.44); p < 0.0001; Figure 5.8 H, I].

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Lean tissue mass at P14

Post-hoc analysis revealed that lean tissue mass (g) was reduced in IUGR + saline compared to control

+ saline pups in sexes combined [-4.35 (-5.68, -3.02); p < 0.0001; Figure 5.9 A], and in males and

females when analysed separately [males: -4.28 (-6.11, -2.45); p < 0.0001; females: -4.42 (-6.37, -

2.47); p < 0.0001; Figure 5.9 B, C]. Lean tissue mass was reduced in IUGR + DITPA compared to

control + DITPA pups when sexes were combined [-3.31 (-4.68, -1.95); p < 0.0001; Figure 5.9 A],

and analysed separately [males: -2.63 (-4.4, -0.85); p = 0.004; females: -3.9 (-6.07, -1.93); p < 0.0001;

Figure 5.9 B, C]. Lean tissue mass was not altered in control or IUGR pups when given DITPA

compared to saline (Figure 5.9 A-C).

Figure 5.8 Bone mineral density (A-C), bone mineral content (D-F), and total bone area (G-I) in male

and female control and IUGR pups at P14 treated with DITPA or saline. Data analysed by linear mixed

modelling and Bonferroni correction. Values presented as Mean ± SEM. * p < 0.05, ** p < 0.01, *** p < 0.001,

**** p<0.0001. Pup numbers: control + saline: n = 34 male, n = 33 female; control + DITPA: n = 33 male, n

= 27 female; IUGR + saline: n = 31 male, n = 18 female; IUGR + DITPA: n = 23 male, n = 24 female.

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Fat mass and percentage body fat at P14

Post-hoc analysis showed that fat mass (g) was reduced in IUGR + saline compared to control + saline

pups in sexes combined [-2.57 (-3.41, -1.74); p < 0.0001; Figure 5.9 D], and in males and females

when analysed separately [males: -2.61 (-3.74, -1.48); p < 0.0001; females: -2.54 (-3.75, - 1.32); p <

0.0001; Figure 5.9 E, F]. Percentage body fat was also reduced in IUGR + saline compared to control

+ saline pups in sexes combined [-2.39 (-4.67, -0.11); = 0.04; Figure 5.9 G] but not in males and

females when analysed separately (Figure 5.9 H, I). Fat mass was reduced in IUGR + DITPA pups

compared to control + DITPA in sexes combined [-1.61 (-2.45, -0.76); p < 0.0001; Figure 5.9 D] and

in males and females when assessed separately [males: -1.29 (-2.39, -0.19); p = 0.022; females: -1.92

(-3.20, -0.64); p = 0.003; Figure 5.9 E, F]. Fat mass and % body fat was not altered in control or

IUGR pups given DITPA treatment compared to saline (Figure 5.9 D-I).

See Appendix 5 for Mean ± SD of body morphometry data.

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5.4.5 Thyroid and liver function

The results of linear mixed modelling analysis showed no interactions between group, treatment, or

sex on circulating FT3, FT4 (both pmol/L), or liver enzymes (ALT, ALP; both IU/L) and cholesterol

(mmol/L) (Type III, p > 0.05). There was a main effect of group on FT4 (F1/123 = 9.9, p = 0.002),

ALT (F1/179 = 12.8, p = 0.001) and ALP (F1/83 = 4.5, 0.04), and an effect of treatment on FT3 (F1/117

=79.1, p < 0.0001), FT4 (F1/123 = 984.2, p < 0.0001), ALT (F1/78 = 7.8, p = 0.007) and ALP (F1/83 =

11.4, p = 0.001). Lastly, there was a main sex effect on FT4 (F1/123 = 9.12, p = 0.003) and ALP (F1/83

= 12.3, p = 0.0001). The human reference ranges for FT3 and FT4 are shown in Table 5.1.

Figure 5.9 Lean tissue mass (A-C), total fat mass (D-F), and percentage (%) body fat (G-I) in male and

female control and IUGR pups at P14 treated with DITPA or saline. Data analysed by linear mixed

modelling and Bonferroni correction. Values presented as Mean ± SEM. * p < 0.05, **, p < 0.001, ****

p<0.0001. Pup numbers: control + saline: n = 34 male, n = 33 female; control + DITPA: n = 33 male, n = 27

female; IUGR + saline: n = 31 male, n = 18 female; IUGR + DITPA: n = 23 male, n = 24 female.

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5.4.5.1 Thyroid function

Free triiodothyronine (FT3)

Post-hoc analysis revealed that there was no difference in FT3 levels (pmol/L) between IUGR + saline

and control + saline pups in sexes combined (Figure 5.10 A), and in males and females when analysed

separately (Figure 5.10 B, C). FT3 was increased in IUGR + DITPA compared to IUGR + saline pups

in sexes combined [5.96 (4.13, 7.78); p < 0.0001; Figure 5.10 A], and in males and females analysed

separately [males: 6.36 (3.87, 8.84); p < 0.0001; females: 5.55 (2.86, 8.24); p < 0.0001; Figure 5.10

B, C]. FT3 was also increased in control + DITPA compared to control + saline pups in sexes

combined [7.56 (5.17, 9.95); p < 0.0001; Figure 5.10 A], and in males and females separately [males:

7.73 (4.45, 11.03); p < 0.0001; females: 7.38 (3.90, 10.86); p < 0.0001; Figure 5.10 B, C]. FT3 levels

were reduced in IUGR + DITPA compared to control + DITPA pups in sexes combined [-2.26 (-4.33,

-0.18); p = 0.34; Figure 5.10 A], and in females only [-3.51 (-6.60, - 0.43); p = 0.026; Figure 5.10 C].

Free thyroxine (FT4)

Post-hoc analysis showed that FT4 levels (pmol/L) were reduced in IUGR + saline compared to

control + saline pups in sexes combined [-2.85 (-4.63, -1.07); p = 0.002; Figure 5.10 D], and in

females [-5.45 (-8.05, -2.85); p < 0.0001; Figure 5.10 F] but not males (Figure 5.10 E) when analysed

separately. FT4 was reduced in IUGR + DITPA compared to IUGR + saline in sexes combined [-

18.68 (- 20.22, - 17.13); p < 0.0001; Figure 5.10 D], and in males and females when analysed

separately [males: -18.31 (-20.38, - 16.23); p < 0.0001; females: - 19.04 (-21.33, -16.76); p < 0.0001;

Figure 5.10 E, F]. FT4 was also reduced in control + DITPA compared to control + saline pups in

sexes combined [-20.46 (-22.39, -18.53); p < 0.0001; Figure 5.10 D], and in males and females when

analysed separately [males: -17.74 (-20.35, -15.12); p < 0.0001; females: -23.19 (-26.02, -20.36); p <

0.0001; Figure 5.10 E, F].

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Table 5. 2 Clinical reference ranges for plasma levels of free triiodothyronine (FT3) and free thyroxine

(FT4) in infants, children and adults. Reference ranges from Monash Pathology, Clayton, Vic, Australia.

Infant (Newborn) Child (< 18 years) Adult ( > 18 years)

FT3 (pmol/L) 5 – 9.4 4.7 – 4.9 3.2 – 6.1

FT4 (pmol/L) 15.3 – 43.6 8.8 – 17.7 8.0 - 16

pmol/L = picomoles per litre.

5.4.5.2 Liver function

Alanine transaminase (ALT) and alkaline phosphatase (ALP)

Post-hoc analysis showed that ALT (IU/L) was decreased in IUGR + saline compared to control +

saline pups when sexes were combined [-9.27 (-14.93, -3.61); p = 0.002; Figure 5.11 A], and in males

and females when analysed separately [males: - 9.18 (-16.58, - 1.79); p = 0.016; females: -9.35 (-

17.78, - 0.93); p = 0.03; Figure 5.11 B, C]. ALT was increased in IUGR + DITPA compared to IUGR

+ saline pups in sexes combined [7.54 (3.05, 12.02); p = 0.001; Figure 5.11 A], and in males and

females analysed separately [males: 8.48 (2.22, 14.73); p = 0.009; females: 6.59 (0.16, 13.02); p =

0.045; Figure 5.11 B, C]. There was no difference in ALT levels between control + DITPA compared

Figure 5.10 FT3 (A-C) and FT4 (D-F) plasma levels in male and female control and IUGR pups at P14

treated with DITPA or saline. Data analysed by linear mixed modelling and Bonferroni correction. Values

presented as Mean ± SEM. pmol/L = picomoles per litre, * p < 0.05, ** p < 0.01, *** p < 0.001, **** p<0.0001.

Pup numbers: control + saline: n = 34 male, n = 33 female; control + DITPA: n = 33 male, n = 27 female;

IUGR + saline: n = 31 male, n = 18 female; IUGR + DITPA: n = 23 male, n = 24 female.

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to control + saline pups, or IUGR + DITPA compared to control + DITPA pups (Figure 5.11 A, B,

C). The human reference ranges for ALT and ALP are shown in Table 5.3.

Post-hoc analysis showed that there was no difference in ALP levels (IU/L) between IUGR + saline

and control + saline pups when sexes were combined (Figure 5.11 D), and when analysed separately

(Figure 5.11 E, F). ALP was increased in IUGR + DITPA compared to IUGR + saline pups in sexes

combined [53.09 (14.24, 91.94); p = 0.008; Figure 5.11 D], and in males [89.81 (35.57, 114.05); p =

0.001; Figure 5.11 E], but not females (Figure 5.11 F). ALP was also increased in control + DITPA

compared to control + saline pups in sexes combined [60.02 (5.65, 114.39); p = 0.03; Figure 5.11 D],

and in males [81.41 (11.38, 151.55); p = 0.023; Figure 5.11 E] but not females (Figure 5.11 F).

Cholesterol

Post-hoc analysis showed there was no difference in cholesterol levels (mmol/L) between any of the

experimental groups when sexes were combined (Figure 5.11 G), or in males and females when

analysed separately (Figure 5.11 H, I).

Dio1:B2M

Two-way ANOVA analysis showed no interaction between group and treatment on the relative

expression of Dio1 to B2M (Dio1:B2M) in the liver. There was no main effect of group in the liver,

but there was an effect of treatment (F1/24 = 5.93, p = 0.02). Post-hoc analysis showed that there was

no difference in Dio1:B2M relative expression in the liver between groups (Figure 5.11).

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Table 5.3 Clinical reference ranges for plasma levels of Alanine transaminase (ALT), alkaline

phosphatase (ALP) and cholesterol in infants, children and adults. Reference ranges from Monash

Pathology, Clayton, Vic, Australia.

Infants (Newborn) Children (< 18 years) Adult (> 18 years)

ALT (IU/L) 5 - 45 - 7 - 56

ALP (IU/L) 80 - 550 120 - 450 30 - 110

Cholesterol

(mmol/L)

0 – 5.5

mmol/L = millimoles per litre, U/L = international units per litre.

Figure 5.11 Serum levels of ALT (A-C), ALP (D-F), and cholesterol (G-I), and relative levels of Dio1 to

B2M (J; male liver only) in male and female control and IUGR pups at P14 treated with DITPA or

saline. Data analysed by linear mixed modelling and Bonferroni correction.2-way ANOVA used to analyse

Dio1:B2M data. Values presented as Mean ± SEM. IU/L = international units per litre, mmol/L millimoles per

litre, * p < 0.05, ** p < 0.01, *** p < 0.001. Pup numbers: control + saline: n = 34 male, n = 33 female; control

+ DITPA: n = 33 male, n = 27 female; IUGR + saline: n = 31 male, n = 18 female; IUGR + DITPA: n = 23

male, n = 24 female. Dio1:B2M analysis: control + saline: n= 6; control + DITPA: n= 6; IUGR + saline: n= 7;

IUGR + DITPA: n= 8

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5.5 Discussion

5.5.1 Overview

This is the first study to examine neonatal growth and wellbeing measures in response to daily, longer-

term DITPA administration from P1 to 13 in an IUGR rat model. The key findings are:

In IUGR + saline compared to control + saline pups there was a significant reduction in (i) body

weight at P1, P7 and P14, crown-to-rump length (CRL), and at P14 a significant reduction in (ii) hip

circumference, (iii) total brain weight, (iv) cerebral hemisphere weight, (v) pons weight, (vi) liver

and kidney weight, (vii) bone mineral density and content, (viii) total bone area, (ix) lean tissue mass,

fat mass, and percentage of body fat, and (x) circulating FT4 and liver enzyme ALT.

In IUGR + DITPA compared to IUGR + saline pups at P14 there was a significant (i) increase in left

and right kidney weight, (ii) increase in plasma FT3 levels, (iii) decrease in plasma FT4 levels,

although these results were due to FT3 cross-reacting with the DITPA assay, and TH homeostatic

feedback mechanism, and (iv) increase in liver enzymes ALT and ALP, although they remained

within safe levels.

In control + DITPA compared to control + saline pups at P14 there was a significant (i) decrease in

body weight of male pups, (ii) decrease in liver weight in female pups, (iii) decrease in pons weight

in males, (iv) increase in bone mineral density in females, (v) increase in plasma FT3 levels, (vi)

decrease in plasma FT4 levels, and (vii) increase in plasma ALP levels. All plasma outcomes were

within safe levels.

Overall, DITPA administration was not harmful to neonatal growth and wellbeing in IUGR rat pups

when assessed at P14, however there is some evidence that it has negative effects on growth, the brain

and liver when administered to control pups.


In the present study body weight at P1, P7 and P14 was significantly reduced, as was liver and kidney

weight at P14 in IUGR compared to control pups. These results are consistent decreased body weight

in IUGR infants (persisting to 1 year of age and into adolescence) (Bernstein et al., 2000, Pena et al.,

1988, Strauss and Dietz, 1998, Martorell et al., 1998), and in the IUGR postnatal rat (Lane et al.,

2001b, Romano et al., 2009, Wlodek et al., 2005, Simmons et al., 1992, McDougall et al., 2017b). As

expected, in the present study liver and kidney weights were reduced as previously reported in IUGR

rats (Cha et al., 1987), and in IUGR infants and children (Naeye, 1965, Hotoura et al., 2005, Schmidt

et al., 2005, Latini et al., 2004, Ladinig et al., 2014).

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In the current study, there was decreased weight gain in male control + DITPA compared to control

+ saline pups at P14. This reduction in growth may be due to a thyrotoxic effect of DITPA when

given to control pups, and this effect is likely masked in the female pups by a higher baseline body

weight of the female control + DITPA pups compared to males at P1 (likely due to random bias) .

These results are consistent with previous studies in humans with euthyroid status, in which DITPA

caused weight loss in adults with congestive heart failure (DITPA administered for 8 weeks, or 6

months at 90 to 180, 270, 360 mg/day; maximum dose of 270mg/day) (Ladenson et al., 2010,

Goldman et al., 2009). Fortunately, DITPA did not affect weight gain in IUGR pups in the present

study. Clinically, DITPA administered to children (aged between 8.5 to 25 months) with congenital

MCT8 mutation (1.8mg/day initially, increased to 30mg/d for 26 to 40 months) improves weight gain

(Verge et al., 2012), however it should be noted that these children had elevated baseline circulating

T3 levels which likely contributed to their weight loss. This is not the case in IUGR foetuses, where

baseline circulating T3 levels are no different from control levels (Thorpe-Beeston et al., 1991). There

are no preclinical studies examining the effects of DITPA treatment in IUGR neonates. However, the

results of the present study are consistent with other rodent studies in MCT8 knockout mice (0.3, 0.6

or 1mg/body weight/day DITPA for 4 days; or 0.3mg/100g/day for 10 days) (Ferrara et al., 2015) (Di

Cosmo, 2009), and in IUGR pups (0.5mg/100g i.p. DITPA daily from P1 – P6) (Azhan, A.,

unpublished thesis, 2019), where body weight was not affected by DITPA administration.

Little is known about the effect of DITPA on kidney and liver growth overall. The present study

found that DITPA treatment in IUGR pups increased right and left kidney weights (left kidney only

when data from both sexes was combined). No difference in circulating plasma levels of T3 has been

reported in IUGR, but circulating T4 levels are decreased compared to controls (Thorpe-Beeston et

al., 1991, Soothill et al., 1992). Hypothyroidism is associated with reduced kidney-to-body mass ratio

in rats, while hyperthyroidism is associated with an increase in kidney-to-body mass ratio (Vargas et

al., 2006). This could explain the finding that DITPA increased kidney weight in IUGR rats if MCT8

is also reduced in the kidneys, as ‘correcting’ a possible lack of TH signalling within the kidneys

could improve their growth trajectory. However there is no evidence that MCT8 is reduced in the

IUGR rat kidney, and further studies examining kidney structure and function in response to IUGR

and DITPA administration are required.

In the present study, the right kidney was more affected by DITPA than the left (effects in left kidney

were evident when male and female data were combined). Rats have slightly heavier right kidneys,

believed to be due to differences in artery morphology (Yoldas and Dayan, 2014) thus perhaps the

ability for the right kidney to process larger amounts of DITPA than the left, contributed to a larger

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effect in the right kidney. In control pups, DITPA significantly decreased liver weight in females

only; at this stage, there is no reason for this sex-specific reduction in liver weight, but does support

the idea that DITPA should not be given to healthy individuals as a pre-emptive measure, as its full

effects on the liver growth and function have not been elucidated. These results suggest that DITPA,

administered from P1 to 13 may be having beneficial result on kidney growth in IUGR in both males

and females, but may have adverse effects in the liver when given to control rats.

5.5.3 Brain weights

This study showed that IUGR + saline compared to control + saline pups, had significantly reduced

total brain weight and weights of the cerebral hemispheres and pons as well as an increased brain-to-

body weight ratio; these findings are in line with our previous rat and guinea pig studies (Tolcos et

al., 2011, Tolcos et al., 2018). An increased brain-to-body weight ratio in IUGR is indicative of foetal

brain sparing that likely occurred in utero. In IUGR infants, the brain is smaller than that of

appropriately grown counterparts (Batalle et al., 2012, Eikenes et al., 2012a), however it may still be

proportionally large compared to body size due to brain sparing. In the present study, there was no

difference in brain weight between IUGR and control pups treated with DITPA or saline, however in

control pups, DITPA administration increased brain-to-body weight ratio and decreased pons weight.

It is currently unclear why this effect was seen only in control males, however as mentioned

previously decreased growth in male control pups could possibly be due to thyrotoxic effects of

DITPA when given to controls, with this effect masked in females as their body weight is heavier

than males at P1. There is currently no data for the effect of DITPA on human brain weight (IUGR

or otherwise), however in IUGR newborn rats treated with DITPA (0.5mg/100g from P1 – P6), brain

weight was not altered, but as in the present study, brain-to-body weight ratio was increased (Azhan,

A., unpublished thesis, 2019). These results suggest that while IUGR reduces brain growth, DITPA

treatment does not worsen these effects.

5.5.4 Body morphometry

Previous IUGR studies report decreased body morphometry measures such as CRL at P7 and P35 in

rats (Azhan, 2019, McDougall et al., 2017b), and in guinea pigs from 52 dg to 8 weeks postnatal age

(Tolcos et al., 2018, Tolcos et al., 2011). The results of the present study are consistent with this

literature, and indicate that CRL is reduced in both male and female IUGR rats (saline treated)

compared to controls at P14; hip circumference was also reduced but in males only. However head

circumference was not different between IUGR and control, saline-treated rats at P14, and may be

due to brain sparing, which results in a proportionally larger brain and head in IUGR pups. IUGR

newborns with these proportions are classified as having asymmetrical IUGR (Godfrey and Barker,

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2001). The presence of brain sparing in IUGR pups in the present study, is supported by the increased

brain-to-body weight ratio in IUGR + saline pups compared to control + saline pups. Body

morphometry measures were not different in IUGR pups treated with DITPA or saline pups in the

present study, or in a previous study using short-term DITPA treatment (0.5mg/100g i.p. daily from

P1 – P6) in IUGR newborn rats (Azhan, A., unpublished thesis, 2019). Unfortunately, previous

human studies using DITPA have focused predominantly on body weight and not body morphometry.

Overall, results of these studies combined, suggest that DITPA does not negatively affect CRL, head

or hip circumference in IUGR.

5.5.5 Body composition (DEXA)

DEXA was used to analyse body composition of pups at P14. The results of DEXA analysis in the

present study revealed that IUGR pups compared to controls had lower bone mineral density, bone

mineral content, total bone area, lean tissue mass, fat mass and percentage of fat mass when assessed

at P14. This is consistent with clinical DEXA data, showing that IUGR foetuses have reduced fat

mass and lean tissue mass (Brown and Hay, 2016, Larciprete et al., 2005). Animal studies also show

reduced DEXA measures in IUGR compared to controls, with reduced total bone area and bone

mineral content in female rats postnatally (Zhao et al., 2013), and decreased fat mass, lean tissue

mass, bone mineral density and content in male rats at P7 (Azhan, A., unpublished thesis, 2019).

DITPA administration in the present study did not alter any of the above mentioned body composition

measures in IUGR rat pups, but did increased bone mineral density in control + DITPA pups

compared to control + saline in sexes combined, and females only when sexes were analysed

separately. It is currently unclear why DITPA produced this effect in control pups, and why no

difference was seen in the males. Notably, DITPA’s thyromimetic properties may elevate bone

turnover in control pups. While TH replacement in hypothyroidism stimulates catch-up growth and

bone maturation, thyrotoxicosis results in increased bone resorption and decreased bone density

(Bassett and Williams, 2003, Harvey et al., 2002, Stevens et al., 2003). DITPA treatment in euthyroid

adults supports this explanation, with increased bone turnover reported in DITPA treated patients

(Ladenson et al., 2010). Importantly in the present study, and our previous study (Azhan, A.,

unpublished thesis, 2019) there was no reduction in the bone mineral density or content in either

control or IUGR pups treated with DITPA. Furthermore, both short- (Azhan, A., Unpublished thesis,

2019) and long-term administration studies (current study) also show that DITPA does not affect lean

tissue mass or fat mass in IUGR newborn rats.

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5.5.6 Blood plasma analysis

Thyroid function

Circulating TH levels are altered in IUGR, with decreased FT3 (Kilby et al., 1998) and FT4 (Soothill

et al., 1992, Thorpe-Beeston et al., 1991, Kush, 2004) levels in IUGR foetuses compared to their

appropriately grown counterparts. In the present study female IUGR pups had decreased FT4 levels

compared to controls (both given saline), but there was no change in males, or in FT3 levels in both

males and females. It is unclear why FT4 levels in males were not affected, as males are considered

more susceptible to the effects of IUGR (Radulescu et al., 2013). Importantly, the TH levels of IUGR

pups in the present study are still within the normal ranges (See Table 5.2).

In the present study, DITPA administration significantly increased FT3 in both IUGR and control

male and female pups, however this result was expected as the plasma assay for FT3 is known to

cross-react with DITPA (Leung et al., 2016). A study using a different assay, which does not cross-

react with DITPA, showed that DITPA (0.3mg/100g body weight/day) normalised plasma T3 to

control levels in MCT8 knockout mice (Ferrara et al., 2015). The findings of the present study

provided evidence that DITPA indeed entered the blood circulation of pups, as DITPA treatment in

both IUGR and control pups significantly decreased FT4 levels. The reason for this decrease is likely

due to homeostatic regulation of TH within the body from a negative feedback system. As previously

discussed, circulating TH levels are biologically maintained at homeostasis, so that when circulating

TH levels are low, the hypothalamus releases TRH which stimulates the pituitary gland to release

TSH, which then signals the thyroid gland to produce more TH (T3 and T4) in the blood stream.

Conversely, if circulating TH levels are high, the hypothalamus and pituitary gland reduce TRH and

TSH levels in order to decrease circulating TH levels. Although DITPA treatment has not been

trialled clinically in IUGR infants, in euthyroid adults with cardiac failure, DITPA caused low

circulating TSH levels (Goldman et al., 2009) likely due to the negative feedback system, but did not

cause signs or symptoms of hypothyroidism or thyrotoxicity. Similarly, in the present study, DITPA

would have signalled the negative feedback system to reduce the production of TSH leading to the

marked decrease in FT4 seen in our assay (see Figure 5.12).

The plasma assay used to measure TSH levels in the present study was not successful (see Section

5.3.4), and limited blood plasma samples meant that the assay could not be repeated. An alternative

approach was to examine Dio1 gene expression in the liver. The Dio1 gene encodes the dominant

enzyme in the liver for activating TH (converting T4 to T3) (Bianco et al., 2002, Kohrle, 1999) and

therefore levels of Dio1 are indicative of the demand for TH in the liver, with low levels of Dio1

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indicating a low demand for TH activation and high levels of Dio1 indicating a higher demand. In the

present study DITPA reduced Dio1 mRNA expression in the liver of pups, irrespective of group

(control or IUGR), and no specific group differences were found using multiple comparisons. A

reduction of Dio1 mRNA in the liver following DITPA administration indicates that less Dio1

enzyme is required as there is a decreased demand for T4 to be activated to T3. Given this lowered

demand for TH activation, it can be assumed that TSH levels are also decreased, as the thyroid gland

is not required to synthesise as much TH, as it is not required. This was a main effect within the data,

however when the specific differences between groups were analysed using post-hoc multiple

comparisons, no differences were found between any of the groups.

Liver function and cholesterol

In the present study, ALT levels, but not ALP or cholesterol levels, were reduced in IUGR compared

to control pups (saline treated), but these were still within the normal range (See Table 5.3). ALT is

present in liver cells only, and serves several purposes, including protein synthesis, storage of iron

and vitamins, bile production to aid digestion and blood detoxification. ALP aids the liver in breaking

down proteins, but is not exclusive to the liver, also being found in bones, intestines and pancreas

(Johnston, 1999). When the liver is damaged or inflamed, these enzymes ‘leak’ out of the cells into

Figure 5.12 Thyroid hormone homeostatic feedback system. Upon disruption of homeostasis, the

hypothalamus either releases TRH (system requires more TH) or does not (system requires less TH). If the

hypothalamus releases TRH, the pituitary gland is stimulated to secrete TSH, which in turn stimulates the

thyroid gland to release more T3 and T4 into the circulation to maintain homeostasis. TRH = thyrotropin-

releasing hormone, TSH = thyroid-stimulating hormone, T3 = triiodothyronine, T4 = thyroxine.

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the blood stream, where they can be measured. In the present study DITPA administration compared

to saline significantly elevated both ALT and ALP (ALP only in males) in IUGR and control pups

when measured at P14, but all levels were still within the normal range. Therefore it is unlikely that

DITPA is benefitting or harming the liver when given in IUGR, and these elevations may just reflect

mild liver cell activation. The mild increase in ALP may also come from increased bone turnover as

a response to DITPA, however DEXA scanning found no evidence of effects on bone in IUGR pups

treated with DITPA compared to saline. Although DITPA corrects excess TH signalling (Di Cosmo,

2009) and a thyrotoxic state in the liver (Ferrara et al., 2015) in MCT8 knockout mice, these mice are

hyperthyroid, and therefore direct comparisons to euthyroid IUGR rats in the present study cannot be

made. Taken together these results suggest that DITPA may slightly activate the liver, but not to the

extent of causing harm. Further studies into the TH state of the liver following DITPA treatment in

IUGR are required.

In regards to cholesterol levels, there are varying results following DITPA administration; in adults

with heart failure, DITPA decreased cholesterol levels following 24 weeks of treatment (Ladenson et

al., 2010). This results differs from the present study, in which the cholesterol levels of IUGR pups

and controls was not altered by DITPA, however the conflicting findings is likely due to the difference

in species, age and dosing regimen.


As mentioned circulating TSH levels, an indirect measure of thyroid function, in the blood of pups

were not measured in the present study. Plasma was allocated for TSH testing, and a standard hospital

plasma assay for TSH was conducted by Monash Pathology. Unfortunately the assay was

unsuccessful, as the specific assay kit was not suitable for rat TSH, and due to limited volume of

plasma collected, there was no remaining plasma to repeat this assay. An alternative approach was to

examine Dio1 activity or gene expression in the liver. This was done using livers of only male animals

(corresponding to those analysed in Chapter 3 and 4), as time did not permit assessment of both males

and females due to University closure in response to the COVID-19 pandemic. Assessment of Dio1

mRNA expression in the liver of female pups will be performed once access to the laboratory

resumes.

5.5.8 Conclusion

In the current study DITPA (0.5mg/100g) or saline (equivalent volume) administered from P1 to 13

to IUGR and control rat pups showed that DITPA does not adversely impact neonatal growth, weights

of brain, kidneys or liver, and body composition following IUGR, despite reducing FT4 levels and

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showing hepatic thyromimetic activity. Although this is promising, before definitive conclusions can

be drawn about the safety profile of DITPA in IUGR newborn infants, the long-term effects of DITPA

on neonatal growth and wellbeing must be examined.

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6 General Discussion

6.1 Overview

IUGR is characterised by foetal growth that is small for GA due to maternal, environmental or genetic

factors, and despite much research, as summarised previously in this thesis, it remains a major clinical

challenge (Murki, 2014). Indeed, IUGR is second to prematurity as the leading cause of perinatal

morbidity and mortality (Abu-Saad and Fraser, 2010, Bhutta et al., 2005), with IUGR babies at an

increased risk of adverse neurodevelopmental sequelae (Geva et al., 2006b), and cerebral palsy

(McIntyre et al., 2013). Deficits in the development of WM (Chase et al., 1972, Eikenes et al., 2012a,

Esteban et al., 2010, Padilla et al., 2014a) and GM (Dubois et al., 2008, Lodygensky et al., 2008,

Tolsa et al., 2004, Benavides-Serralde et al., 2009) as well as delayed brain maturation are thought to

underlie neurodevelopmental impairments associated with IUGR. It is now accepted that disrupted

and delayed myelination is often present in the IUGR infant’s brain (Chase et al., 1972), and this

altered WM development can persist into adulthood (Padilla et al., 2014a, Esteban et al., 2010,

Eikenes et al., 2012b). In the CNS, OLs are essential for making myelin, and undergo a lineage

progression from progenitors to mature myelinating OLs, driven in part by TH and cerebral TH

signalling. The relationship between TH signalling and OL maturation has been the focus of this

thesis.

Previously, our group (Azhan, A., unpublished thesis, 2019), and others (Chan et al., 2014) have

found that the TH transporter MCT8, which exclusively transports TH into cells in the brain,

including into OLs themselves, is down-regulated in the brain of IUGR human foetuses and newborn

rats. This suggests that a deficit in cellular transport of TH could underlie myelination impairments

observed in IUGR. This proposal is supported by evidence of reduced myelination in children with a

congenital MCT8 mutation (Lopez-Espindola et al., 2014). Our group has previously examined the

impact of a short term dosing regimen of DITPA (P1 to P6) in IUGR and control rats, and found that

by P7, myelination was restored in the brains of IUGR rats (Azhan, A., unpublished thesis, 2019).

However to extend this research, it was necessary to study a longer dosing regimen of DITPA; i.e.,

one which was more relevant to the clinical scenario of an IUGR preterm neonate receiving DITPA

until term equivalent age. If severe or worsening IUGR is detected in utero, the baby is often delivered

preterm therefore ideally DITPA treatment would be given to these preterm IUGR babies, until the

time at which MCT8 levels normalise in their brain. Although data related to the temporal expression

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of MCT8 in the IUGR human brain is unavailable, previous findings from our group report that MCT8

levels are reduced in the IUGR rat brain at P7, and these normalise by P14. Therefore in the present

study, the effects of DITPA administration in IUGR rats were examined from P1 to P13, a time in rat

brain development considered to be equivalent to that of a human baby from 23 to 40 weeks GA

(Semple et al., 2013). Thus, the experiments summarised in Chapters 3 to 5 of this thesis are reflective

of what is likely to occur in the clinical setting where an IUGR baby is delivered preterm and then

treated with DITPA until term equivalent age. The studies were designed to examine the impact of

daily DITPA treatment over 1 to13 days after birth (P1 to P13), on myelination and OL development

in the brain. As discussed previously (Chapter 1 to Chapter 5), DITPA is a TH analogue that does not

require MCT8 to enter cells in the brain. A focus was placed on examining regions of the brain known

to be highly vulnerable to prenatal insults like IUGR, including the cerebral cortex, corpus callosum,

hippocampus, and the cerebellum (Padilla et al., 2011, Padilla et al., 2014b, Egana-Ugrinovic et al.,

2014, Lodygensky et al., 2008). The aims of the study were to determine if longer-term DITPA

treatment in IUGR rat pups would restore myelination and promote OL maturation, without causing

injury or inflammation in the brain.

The major findings are that DITPA, when administered to IUGR pups, promoted MBP-IR in the

cortical layer VI and fimbria, and increased OL cell density (Olig2) in the corpus callosum, but also

decreased myelin protein (PLP-IR) in the corpus callosum and external capsule at P14 compared to

IUGR pups administered saline. In the IUGR cerebellum, DITPA had no effect on MBP-IR and PLP-

IR coverage or Olig2-IR cell density, but did increase the linear density of Purkinje cells (cells/mm),

the main motor output cell of the cerebellum in the early developing lobules. DITPA treatment did

not cause injury or inflammation (assessed via Iba1-IR microglia and GFAP-IR astrocytes) in any

cerebral structure, or in the cerebellum, albeit for a small but significant increase in the density of

Iba1-IR microglia in late developing cerebellar lobules. Importantly, DITPA did not adversely impact

neonatal growth, brain or organ weights, or body composition in IUGR pups. The present study found

that DITPA elevated plasma FT4 levels in IUGR pups, however this was likely due to a TH feedback

mechanism; importantly FT4 levels remained within normal ranges and overall thyroid function was

normal. DITPA also elevated liver enzymes ALT and ALP, however these also remained within

normal ranges, suggesting that DITPA is unlikely to alter liver function significantly when given to

IUGR infants. Results from each study, as well as the strengths and weaknesses of each study have

been discussed in Chapters 3 to 5. The following discussion will focus on how the findings presented

in this thesis contribute to our understanding of DITPA as a therapy for brain injury following IUGR.

Future directions of this research will also be discussed.

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6.2 Does DITPA promote myelination in the cerebrum and therefore

benefit the IUGR brain?

The impact of DITPA treatment on the IUGR brain is currently not well understood, however in the

present study it was expected that DITPA would have a pro-myelinating effect in the brains of IUGR

rats, as previous research by our group has shown that daily DITPA administration from P1 to P6

restored myelination by P7 in the external capsule of the IUGR rat brain compared to saline treatment

(Azhan, A., unpublished thesis, 2019). As previously discussed in this thesis, IUGR is reported to

elevate genes which repress OL maturation (WNT, NOTCH & BMP4) and supress those which

promote OL maturation and myelination (SOX10 & Myrf; Azhan, A., PhD thesis, Monash

University, 2019). It is not yet clear what triggers these genetic changes that result in impaired

myelination, however there is proof that hypoxic-ischaemic insults in-utero, like those occurring

during IUGR may play a part through oxidative stress (French et al., 2009, Reid et al., 2012, Back et

al., 2002). Prenatal hypoxia has been shown to increase WNT signalling in the brain, with elevated

Axin2 levels (a transcriptional target of the WNT signalling pathway) found within OPCs (Fancy et

al., 2011) as well as the overexpression of NOTCH’s downstream target gene Hes5 (Wang et al.,

1998, Zhang et al., 2009). DITPA has also been shown to up-regulate MBP gene expression and to

promote human OL maturation in vitro (Lee et al., 2017). Raising the question of whether DITPA is

impacting the altered gene expression seen in IUGR. However, this thesis showed that DITPA had a

pro-myelinating effect in the cerebrum, irrespective of experimental group, with increased MBP-IR

seen in cortical layer VI and the fimbria in both IUGR and control pups treated with DITPA,

compared to treatment with saline. Of interest, the length of MBP-IR myelinated processes extending

through the cortical layers in these animals were shorter in pups treated with DITPA compared to

saline, and this was true for both IUGR and control pups. These results suggest that although DITPA

increases MBP-IR in cortical layer VI, there may be a delay in the propagation of MBP along cortical

processes, which was visible at P14. Once myelin protein is synthesised in an OL cell body, it

migrates distally into the OL processes where it wraps around neuronal axons (Pedraza, 1997). One

explanation for why MBP-IR does not extend as far along the processes, may be that DITPA is

slowing the MBP migration, meaning that overall, myelination is delayed. The rate of formation of

mature myelin throughout axonal processes is clearly something that should be investigated further.

Another possible explanation for the delayed progression of MBP along cortical processes, as well as

the increased density of MBP-IR seen in cortical layer VI, is that DITPA is causing a ‘bottleneck’

effect; i.e., blocking MBP from travelling properly from the OL body along the cortical processes.

Cortical neuronal projections are fundamental for output and integration of information within the

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brain, and myelination of these projections is essential for optimal brain function. Therefore future

studies should investigate DITPA’s effect on MBP migration along cortical axons at various time

points prior to, and after P14 to better understand if the delayed MBP migration along processes is

due to a delay migration delay or a blockage, and to importantly establish whether myelination of

these processes eventually normalises.

In the present study DITPA treatment had no effect on Olig2-IR or APC-IR cell density, or areal

coverage of MBP-IR or PLP-IR in the IUGR cerebellum. This was surprising, as in the rat the

cerebellum undergoes a rapid period of growth in the first 10 days of life (Dobbing and Sands, 1979),

and cerebellar myelination occurs from P1 – P45 (Hamano et al., 1998). Therefore it was expected

that DITPA administered during this window would impact myelination, and restore any deficits in

cerebellar growth or structure. Although other studies have determined that MCT8 is deficient in the

cortical plate of human IUGR foetuses (Chan et al., 2014), and in cerebral WM of IUGR postnatal

rats (Azhan, A., unpublished thesis, 2019), none have investigated if IUGR renders the cerebellum

deficient in MCT8 expression. It may be the case that MCT8 expression in the cerebellum is not

affected by IUGR, and therefore DITPA treatment would have no benefit, as observed in the current

study. Future studies should examine the expression of MCT8 protein or mRNA in the cerebellum of

IUGR and control pups using immunohistochemical and molecular techniques.

DITPA treatment increased the density of Iba1-IR microglia in late developing cerebellar lobules in

IUGR pups, possibly suggesting an inflammatory response. Microglia are key immune defence cells

in the CNS, however microglia also have other important roles in brain development such as

regulating the migration of neuronal precursors (Erblich et al., 2011), synaptic pruning, modulating

neuronal function (Cserep et al., 2020), and regulating OL differentiation (Aarum et al., 2003). There

is good evidence that microglia are essential for myelination by phagocytic removal of cellular debris

which would otherwise inhibit the recruitment and maturation of OLs and ensheathing of neuronal

axons (Kotter et al., 2006, Saederup et al., 2010). In the present study the thickness of tissue sections

(8µm), prevented the definitive determination of reactive (amoeboid) versus resting (ramified)

microglia, as microglial processes extend in all directions over many microns. Ramified microglia in

the CNS are characterised by long extended processes, and are crucial for neuronal health, as well as

clearing dead cells, and pruning and remodelling neuronal processes in the foetal brain (Schafer et

al., 2012, Walker et al., 2014). Reactive microglia on the other hand have retracted processes and are

classically associated with an inflammatory response to infection or injury (Graeber et al., 2011).

Without knowing the state of the microglia in the late cerebellar lobules in the present study, there is

insufficient information to comment on the nature of the increase in microglial density observed in

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response to DITPA. Future studies should therefore investigate microglial morphology in response

to DITPA treatment using thicker tissue sections to image the full depth of the section.

In this study DITPA administration increased the linear density (cells/mm) of Purkinje cells in IUGR

(and control) pups, in the early developing lobules of the cerebellum. Whether DITPA promotes

Purkinje cell proliferation, or inhibits the process of natural Purkinje cell death in the rat cerebellum

cannot be determined from the findings of the current study. It is also unclear why this result was

only seen in the early developing lobules, or what the functional outcomes of this change would be.

Due to the design of this study, behavioural testing at P14 was not possible, however it would be

relevant for future studies to examine the effect of DITPA administration on behavioural functions,

especially motor coordination, as the cerebellum and Purkinje cells are essential for this function

(Wulff et al., 2009).

6.3 Is DITPA only beneficial when cerebral MCT8 is reduced?

In human pregnancies, if IUGR is detected in utero the only current intervention is to deliver the baby

preterm to remove it from the insufficient intrauterine environment; thus, many IUGR babies are born

preterm (between 24 to 40 weeks GA). In a clinical setting, DITPA therapy would ideally be given

to an IUGR baby until a time at which MCT8 levels normalise, when DITPA would no longer be

required as TH would be able to enter cells via the MCT8 carrier. We know from a previous study in

rats, that MCT8 mRNA levels are reduced in the IUGR rat brain at P7 compared to controls but are

normalised to control values by P14 (Azhan, A., unpublished thesis, 2019), an age equivalent to brain

development in a human baby at term age (~40 weeks GA) (Semple et al., 2013); this knowledge

provided the current study with an understanding of an appropriate duration of DITPA administration

(i.e. P1 to P13). Although we know that MCT8 levels in the brains of IUGR rats normalise by P14

(Azhan, A., unpublished thesis, 2019), we do not know at exactly which point in development

between P7 and P14 this normalisation occurs, and whether an equivalent recovery also occurs in

human babies. IUGR post-mortem studies in human infants are rare, and therefore ontogeny studies

in animals would be valuable for understanding the spatial and temporal profile of MCT8 expression

following IUGR, and in determining the timing of MCT8 recovery. It is quite possible that MCT8

levels in the brains of the IUGR rat pups reported here normalised before P14, and thus after that

point DITPA would serve no purpose other than to supply excess TH-analogue to a system which,

potentially, no longer needed it. This in turn could have negative implications for brain development

after IUGR such as the increased density of microglia seen in the cerebellum as mentioned above, or

perhaps causing the decrease in PLP-IR observed in the corpus callosum and external capsule of

IUGR pups. In the present study, when DITPA was administered to control pups, there were some

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adverse effects which will be discussed in the next section, supporting the proposal that DITPA

should not be given in a system with normal TH signalling. Therefore, it is essential that we

understand precisely when MCT8 levels normalise in the IUGR human infant, so that we can avoid

using a therapy at a time when it may no longer be beneficial, and may in fact be detrimental.

6.4 Should DITPA only be used in cases of confirmed IUGR?

Clinical studies show that when DITPA is administered to euthyroid adults, there are undesirable

effects such as increased bone turnover and reduced cholesterol levels (Ladenson et al., 2010). The

current study found that when DITPA was given to control pups, it reduced PLP-IR in the

hippocampus, external capsule and the fimbria, decreased density of the overall population of OLs

(i.e. Olig2-IR OLs) in cortical layer VI, and reduced the proportion of mature APC-IR OLs in the

overall Olig2-IR OL population; this indicates that in control animals DITPA may impair maturation

of the OL lineage. Given that the density of mature OL was reduced, it was surprising to find that

DITPA increased MBP-IR in cortical layer VI of control pups, possibly suggesting that the existing

OLs accelerate the process of myelination, perhaps as a compensation for the reduced number of

mature OLs. In the cerebellum, control pups treated with DITPA had a 15% reduction in the linear

density of BG fibres in the late developing lobules, compared to pups treated with saline. BG fibres

act as scaffolds along which neurons migrate from the outer proliferative zone of the cerebellum to

the internal granular layer, and therefore fewer BG fibres may indicate disruption to neuronal

migration in the cerebellum. However this was not seen in IUGR pups treated with DITPA compared

saline in this study. The increased BG fibre density seen in control pups may be the effect of DITPA

being administered when the neonatal thyroid status is essentially normal.

At P14, DITPA administration resulted in an 8% decrease in body weight of males, a decrease in the

weight of the pons in males, and an increase in bone mineral density in females; this was not seen in

a previous study when DITPA was given for a shorter duration (P1 to P6) (Azhan, A., unpublished

thesis, 2019). The reduction in body weight observed in males may be due to a thyrotoxic effect of

DITPA when given under euthyroid circumstances, and is likely masked in females due to a higher

baseline body weight at P1. It is unclear why the pons was affected, in the absence of an effect in

other brain regions, raising the question of whether the pons is more vulnerable than the rest of the

brain to the effects of DITPA, or TH treatment in general. The pons is the site of deep cerebellar

nuclei (Brodal and Bjaalie, 1992) and the superior, mid and inferior cerebellar peduncles which are

fibre tracts responsible for connecting the cerebellum with the rest of the CNS (Glickstein and Doron,

2008). Given this connection between the cerebellum and the pons, it is entirely possible that the

responses to DITPA seen in the cerebellum in this study, including increased Purkinje cell density

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(cells/mm) in the early developing lobules, increase microglial density (cells/mm2) and increased in

BG fibres (cells/mm) in the late lobules are related to changes in the pons however this requires

further investigation.

The increased bone density observed in control female pups treated with DITPA compared to saline,

is in line with the finding of increased bone turnover measures in humans (Ladenson et al., 2010) and

should be carefully considered. It is not clear why this was not seen in males, however one possible

explanation is that the hormonal differences between the sexes may be playing a part. The results of

the current study, as well as those in euthyroid adults (Goldman et al., 2009, Ladenson et al., 2010),

suggest that DITPA should only be given to individuals who are confirmed as IUGR, and not as a

prophylactic treatment when IUGR is suspected such as when an infant is small for gestational age,

as adding DITPA to a euthyroid system may cause more harm than good.

6.5 Future directions – clinical administration of DITPA

In the current study DITPA was administrated to rat pups daily using intraperitoneal injections.

Although this is an easy and efficient route of DITPA administration to neonatal rats, it is clearly not

an appropriate delivery route for IUGR infants. Intravenous delivery of drugs is commonly used in

neonates, and is suitable for DITPA administration. Oral administration of drugs in neonates is less

common, and given that the gastrointestinal system in IUGR babies is immature (Bozzetti et al., 2012,

Nicholl et al., 2008), and may also be under physiological stress (e.g., if necrotising enterocolitis is

present (Bernstein et al., 2000)), this may not be ideal as DITPA may not be adequately and rapidly

absorbed; admittedly, there is no data to confirm or refute this proposition. Also, the pain and

discomfort associated with replacement of intravenous catheters necessary for long-term

administration of DITPA should be avoided. The most effective and non-invasive route of

administering DITPA to IUGR babies requires further investigation, with possible administration

options including direct pulmonary delivery via aerosolisation, or by using nanoemulsions.

Pulmonary drug delivery involves drugs being aerosolised via propellants or pressure and inhaled

directly into the patient’s lungs; many forms, including nebulisers, metered dose inhalers and dry

powder inhalers are already available. A nebuliser administers aerosol through a mask, and does not

require a specific inhalation technique, making it ideal for use in infants, unlike metered dose and dry

powder inhalers which do require specific inhalation techniques. Another promising avenue of

administration is via nanoemulsions, which consist of two immiscible liquids combined into droplets

of approximately 100 – 300 nm in size and stabilised by surfactants (Anton et al., 2008, Comfort et

al., 2015), to act as a carrier for the drug. Preclinical trials show that when administered intranasally,

nanoemulsions carrying drugs can bypass the blood-brain barrier and reach the brain via the olfactory

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and trigeminal nerves in the upper nasal cavity (Chapman et al., 2013, Hanson and Frey, 2008,

Bonferoni et al., 2019). This method is non-invasive, does not require patient coordination, displays

rapid drug onset and minimises systemic exposure making it an ideal candidate. Although these are

all possible routes for administering DITPA to IUGR babies, the pharmacodynamics and

pharmacokinetics of DITPA used for each of these methods would need to be considered.

6.6 Conclusion

In summary, this thesis has for the first time demonstrated that the TH analogue DITPA, which enters

cells independently of the MCT8 transporter, when administered for a clinically relevant duration in

IUGR pups (P1 to P13; 0.5mg/100g/day i.p.), promotes myelination (as indicated by increased MBP-

IR) in cortical layer VI, increases the density of OLs in the corpus callosum, does not cause injury or

inflammation to the brain except for a possible minor inflammatory response in the cerebellum, and

has no serious negative off-target effects on neonatal growth or wellbeing. The rat cerebellar WM is

not affected by IUGR at P14, and may therefore serve for interspecies comparisons in order to

elucidate pathways of injury and resistance against IUGR. DITPA treatment in individuals with

normal TH levels (or normal cerebral TH signalling) should be avoided, as this thesis reports

unfavourable results in control rats given DITPA, likely the result of excess TH being administered

into an otherwise normal system. Genes which regulate OL development and are affected in IUGR

may be impacted by DITPA administration, improving outcomes. Future studies should therefore

aim to further examine the neuroprotective potential of DITPA across species. The spatial and

temporal expression profile of MCT8 in the IUGR human and animal brain should also be explored,

as this will provide greater understanding of the point at which MCT8 levels normalise, to better tailor

treatment. This information would greatly aid in the further development and translation of therapies

like DITPA, which may compensate for MCT8 deficiency in IUGR.

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8, e82713.

ZOELLER, R. T. & ROVET, J. 2004. Timing of thyroid hormone action in the developing brain: clinical

observations and experimental findings. J Neuroendocrinol, 16, 809-18.

Appendix 1

DITPA preparation for P1 to P13 rat injections

1. Make up 0.1M NaOH:

a. Dissolve 2g of NaOH in 500mL of H2O

b. Autoclave solution

c. Place into 1.5mL/2mL Eppendorf when ready to use

d. Use a syringe attached to a 40μm filter (Trajan Scientific, Vic, Australia) then draw up 0.1M

NaOH and transfer into Eppendorf

e. Label tubes 0.1M NaOH

2. 0.9% sterile Saline (FreeFelx, Code K690521)

a. Transfer the saline into multiple PCR tubes (placed in Eppendorf tube) using a syringe

b. Label 0.9% saline

3. To make 25mg/6000μL of DITPA

a. Measure 25mg of DITPA into Falcon tube or large 5mL Eppendorf

b. Slowly add 1mL of 0.1M NaOH, vortex each time

c. Add 4mL of 0.1M NaOH to 2mL of 0.9% Saline

4. Once solution has dissolved

a. Transfer the solution into multiple PCR tubes (placed in Eppendorf tube) using a syringe

b. Label 25mg/6000μL DITPA

The author acknowledges the scientific and technical assistance of Ms Aminath Azhan, Monash University.

Product: 3-[4-(4-hydroxyphenoxy)-3, 5-diiodophenyl] propionic acid (DITPA)

Company: Ryan Scientific Inc.,

Catalogue Number: ALBB-014981

https://www.ryansci.com/products/7671341/view

Appendix 2

Optic nerve processing for Transmission Electron Microscopy

Protocol provided by Sarah Ellis, Head of Centre.

1. Following primary fixation at RMIT (Section 2.7.4 above), optic nerves were rinsed in 0.1M

sodium cacodylate buffer.

2. Following this, optic nerves were post-fixed in 1% osmium tetroxide, 1.5% potassium ferrocyanide

in 0.1M sodium cacodylate buffer.

3. Optic nerves were subsequently rinsed in distilled water then dehydrated through a graded series

of alcohols before embedding in Spurrs Resin (Spurr 1969) according to standard electron microscopy

protocol.

References:

Spurr, A.R. 1969. A low-viscosity epoxy resin embedding medium for electron microscopy. J.

Ultrastructure. Res. 26: 31

Appendix 3

Appendix 3, Table 3. 1 Two-way ANOVA results of MBP-IR analysis, supplementary to Chapter 3,

Section 3.7.1. The effect of group (control or IUGR) and treatment (saline or DITPA) on MBP-IR in brain

regions within the cerebral hemispheres is shown. Data are presented as M ± SEM, and values represent

percentage of area covered by MBP-IR (% AC), except for cortical projection, which is presented as a length

ratio (projection length (mm)/cortex width (mm)).

Control IUGR

MBP-IR Saline n= 8 DITPA n= 7 Saline n= 8 DITPA n= 8

Cortical layer VI

31.67 ± 0.87 43.15 ± 1.56 27.94 ± 0.95 36.33 ± 1.45

Cortical projections 60.03 ± 0.97 56.19 ± 2.13 56.67 ± 1.17 50.44 ± 2.20

Corpus callosum 55.16 ± 2.37 58.71 ± 5.04 53.64 ± 3.43 46.47 ± 4.06

External capsule 70.03 ± 1.88 76.73 ± 2.35 74.03 ± 2.31 73.89 ± 1.88

Hippocampus CA1 13.81 ± 0.82 14.48 ± 0.84 10.03 ± 0.57 8.00 ± 0.50

Hippocampus CA3 17.00 ± 0.90 18.47 ± 0.77 13.39 ± 0.86 13.80 ± 1.14

Fimbria 41.73 ± 0.92 75.55 ± 3.33 40.02 ± 1.18 74.07 ± 3.83

Appendix 3, Table 3. 2 Two-way ANOVA results of PLP-IR analysis, supplementary to Chapter 3,

Section 3.7.1. The effect of group (control or IUGR) and treatment (saline or DITPA) on PLP-IR in brain

regions within the cerebral hemispheres is shown. Data are presented as M ± SEM, and values represent

percentage of area covered by PLP-IR (% AC), except for cortical projection, which is presented as a length

ratio (projection length (mm)/cortex width (mm)).

Control IUGR

PLP-IR Saline n= 8 DITPA n= 7 Saline n= 8 DITPA n= 8

Cortical layer VI

20.96 ± 1.63 16.01 ± 1.78 7.91 ± 0.85 6.79 ± 1.02

Cortical

projections

37.42 ± 6.22 28.46 ± 5.52 32.38 ± 5.69 26.96 ± 5.33

Corpus Callosum

28.76 ± 4.37 20.68 ± 3.40 19.55 ± 3.72 7.09 ± 0.83

External capsule

43.56 ± 1.36 34.87 ± 1.41 30.44 ± 1.36 22.82 ± 1.44

Hippocampus

CA1

11.78 ± 2.75 5.63 ± 1.36 2.08 ± 0.37 1.75 ± 0.41

Hippocampus

CA3

13.43 ± 1.19 13.97 ± 3.08 5.74 ± 1.06 7.88 ± 1.62

Fimbria 18.28 ± 2.18 14.04 ± 2.43 6.99 ± 1.12 7.41 ± 1.01

Appendix 3, Table 3. 3 Two-way ANOVA results of Olig2–IR analysis, supplementary to Chapter 3,

Section 3.7.1. The effect of group (control or IUGR) and treatment (saline or DITPA) on Olig2-IR cell density

in brain regions within the cerebral hemispheres is shown. Data are presented as M ± SEM, and values

represent cell density (cells/mm2).

Control IUGR

Olig2-IR cell

density

Saline n= 8 DITPA n= 7 Saline n= 8 DITPA n= 8

Cortical layer VI 901.97 ± 21.22 805.03 ± 21.58 791.90 ± 22.18 838.15 ± 20.54

Corpus callosum 2006.60 ± 29.02 2251.19 ± 134.83 1718.06 ± 113.59 2252.08 ± 125.45

Hippocampus

CA1

1346.35 ± 115.35 1331.55 ± 59.58 1243.75 ± 70.86 1325.00 ± 56.45

Hippocampus

CA3

1444.27 ± 90.12 1338.69 ± 52.84 1332.29 ± 54.80 1454.69 ± 69.04

Fimbria 3186. 46 ± 168.71 2942.86 ± 163.14 2432.29 ± 183.12 3030.21 ± 255.03

Appendix 3, Table 3. 4 Two-way ANOVA results of APC–IR analysis, supplementary to Chapter 3,

Section 3.7.1. The effect of group (control or IUGR) and treatment (saline or DITPA) on APC-IR cell density

in brain regions within the cerebral hemispheres is shown. Data are presented as M ± SEM, and values

represent cell density (cells/mm2).

Control IUGR

APC-IR cell

density



Corpus callosum 958.33 ± 44.86 857.69 ± 90.42 535.71 ± 32.82 689.68 ± 42.16

Hippocampus

CA1

663.02 ± 58.63 419.44 ± 148.56 287.50 ± 51.07 397.02 ± 49.93

Hippocampus

CA3

715.63 ± 44.65 518.06 ± 173.80 337.50 ± 67.53 471.43 ± 58.63

Fimbria 1833.33 ± 125.40 1152.08 ± 380.16 975.00 ± 185.67 1272.02 ± 192.60

Appendix 3, Table 3. 5 Two-way ANOVA results of APC-IR: Olig2-IR analysis, supplementary to

Chapter 3, Section 3.7.1. The effect of group (control or IUGR) and treatment (saline or DITPA) on the

percentage of mature APC-IR OLs within the overall Olig2-IR OL population, in brain regions within the

cerebral hemispheres is shown. Data are presented as M ± SEM, and values represent percentage of mature

APC-IR OLs within the overall Olig2-IR OL population (%).

Control IUGR

APC-IR: Olig2-

IR (%)



Corpus callosum 0.53 ± 0.03 0.52 ± 0.05 0.38 ± 0.03 0.53 ± 0.07

Hippocampus

CA1

51.05 ± 5.68 41.65 ± 12.32 24.06 ± 4.57 26.33 ± 5.06

Hippocampus

CA3

50.25 ± 3.18 36.28 ± 11.53 24.78 ± 4.61 27.70 ± 4.93

Fimbria 57.72 ± 2.94 28.08 ± 10.55 28.21 ± 5.38 35.91 ± 5.96

Appendix 4

Appendix 4, Table 4. 1 Two-way ANOVA results of cerebellar morphology measurements using H&E

staining supplementary to Chapter 4 Section 4.4.1: Areas are in mm2, width is in mm, ratios are % of the

total cross sectional area. Data are presented as M ± SEM.

Control IUGR

Saline n = 8 DITPA n = 7 Saline n = 8 DITPA n = 8

TCA 12.05 ± 0.27 11.47 ± 0.55 10.64 ± 0.41 10.41 ± 0.27

ML area 5.77 ± 0.20 5.44 ± 0.26 4.88 ± 0.13 4.86 ± 0.16

IGL area 5.58 ± 0.40 5.05 ± 0.33 4.22 ± 0.29 4.45 ± 0.22

WM area 1.38 ± 0.06 1.16 ± 0.05 1.21 ± 0.12 1.04 ± 0.07

GM area 7.00 ± 0.45 6.64 ± 0.28 6.64 ± 1.08 5.10 ± 0.64

ML : TCA 47.84 ± 1.04 47.44 ± 0.87 47.30 ± 1.49 46.71 ± 0.82

IGL : TCA 46.38 ± 3.45 34.99 ± 1.78 40.62 ± 0.67 43.25 ± 2.24

WM : TCA 11.50 ± 0.51 10.29 ± 0.70 11.63 ± 0.75 10.13 ± 0.69

GM : TCA 58.33 ± 4.19 58.12 ± 1.77 64.46 ± 10.64 49.67 ± 6.60

ML width 0.10 ± 0.002 0.11 ± 0.002 0.10 ± 0.003 0.10 ± 0.003

GM = grey matter, IGL = internal granular layer, ML = molecular layer, TCA = total cerebellar cross-sectional-

area, WM = white matter

Appendix 4, Table 4. 2 Two-way ANOVA results of immunohistochemical analysis in the cerebellum,

supplementary to Chapter 4 Section 4.4.2. Data are presented as M ± SEM. Values represent percentage of

area covered by MBP-IR (% AC), cell density (cells/mm2), and linear cell density (cells/mm).

Control IUGR

Saline n = 8 DITPA n = 7 Saline n = 8 DITPA n = 8

MBP-IR

DWM

Lobules

74.30 ± 4.82

53.19 ± 5.22

74.32 ± 3.39

49.36 ± 5.49

73.88 ± 2.68

42.79 ± 6.01

72.64 ± 4.64

52.72 ± 3.31

Olig2-IR

DWM

Lobules

2730.40 ± 322.19

2349.08 ± 110.27

3671.52 ± 503.99

2734. 84 ± 250.68

3656.66 ± 511.57

2896.50 ± 198.93

3777.29 ± 400.30

3004.68 ± 341.14

Iba1-IR

DWM

Early lobules

Late lobules

471.43 ± 47.38

515.65 ± 62.80

463.08 ± 72.87

417.86 ± 62.13

301.00 ± 87.45

453.61 ± 98.87

346.43 ± 48.00

588.73 ± 104.30

406.22 ± 98.50

434.38 ± 33.05

530.00 ± 106.51

957.55 ± 131.47

GFAP-IR BG

Early lobules

Late lobules

165.16 ± 4.53

156.41 ± 2.41

147.68 ± 5.46

132.32 ± 4.10

177.34 ± 4.09

176.56 ± 5.52

183.44 ± 4.32

184.53 ± 5.50

GFAP-IR astrocytes

DWM

Lobules combined

Early lobules

Late lobules

18.76 ± 1.58

42.27 ± 1.72

43.13 ± 4.73

41.42 ± 8.69

23.27 ± 4.69

36.26 ± 1.69

36.83 ± 2.62

35.69 ± 5.80

25.66 ± 3.66

47.27 ± 3.16

46.12 ± 3.69

48.41 ± 15.19

27.86 ± 3.06

46.66 ± 2.78

47.22 ± 3.60

46.10 ± 12.70

Calbindin

PC somal area

(lobules combined)

PC areal density

(lobules combined)

PC linear density

Lobules combined

Early lobules

Late lobules

293.68 ± 6.46

1173.03 ± 76.03

34.84 ± 1.07

35.00 ± 1.16

34.69 ± 1.73

309.90 ± 13.01

1295.08 ± 101.23

38.39 ± 30.2

43.57 ± 2.98

33.21 ± 3.44

290.26 ± 11.52

1275.64 ± 84.03

34.53 ± 0.97

33.75 ± 1.06

35.31 ± 1.60

294.62 ± 6.19

1365.12 ± 59.91

40.47 ± 1.55

42.81 ± 1.53

38.13 ± 2.75

DWM = deep white matter, GFAP = glial fibrillary acidic protein, GM = grey matter, Iba1 = ionized calcium-

binding molecule 1, IGL = internal granular layer, MBP = myelin basic protein, ML = molecular layer, Olig2

= oligodendrocyte transcription factor-2, PC = Purkinje cell, TCA = total cerebellar cross-sectional-area, WM

= white matter

Appendix 5

Control IUGR

Sex

Saline n = 23 males

n = 24 females

DITPA n = 31 males

n = 18 females

Saline n = 33 males

n = 27 females

DITPA n = 34 males

n = 33 females

P1 (g) M 6.68 ± 0.61 6.78 ± 0.66 4.84 ± 0.71 4.98 ± 0.57

F 6.31 ± 0.55 6.88 ± 0.75 4.79 ± 0.73 4.78 ± 0.62

P7 (g) M 17.06 ± 1.42 16.38 ± 2.24 11.59 ± 2.28 12.29 ± 2.42

F 16.58 ± 1.43 16.83 ± 2.58 11.97 ± 2.56 11.96 ± 2.19

P14 (g) M 35.45 ± 1.85 32.69 ± 3.53 27.79 ± 4.28 27.90 ± 3.48

F 34.86 ± 2.23 33.06 ± 4.71 28.48 ± 4.66 27.41 ± 4.85

CRL (mm) M 80.96 ± 8.26 81.29 ± 6.86 76.48 ± 8.77 79.79 ± 8.74

F 80.43 ± 9.96 79.50 ± 9.10 78.47 ± 16.37 77.31 ± 6.98

Head Circ. (mm) M 62.96 ± 5.24 64.38 ± 4.12 61.00 ± 5.37 60.25 ± 6.22

F 60.49 ± 13.34 63.00 ± 4.96 61.35 ± 3.70 60.24 ± 4.06

Hip Circ. (mm) M 76.48 ± 6.81 73.04 ± 6.26 68.86 ± 5.17 68.82 ± 6.07

F 71.28 ± 16.87 73.44 ± 8.43 69.92 ± 5.26 70.48 ± 4.58

Liver (g) M 0.91 ± 0.21 0.86 ± 0.28 0.68 ± 0.19 0.61 ± 0.12

F 1.01 ± 0.22 0.85 ± 0.30 0.70 ± 0.20 0.66 ± 0.17

L Kidney (g) M 0.20 ± 0.03 0.20 ± 0.02 0.14 ± 0.03 0.16 ± 0.04

F 0.20 ± 0.03 0.22 ± 0.05 0.16 ± 0.04 0.18 ± 0.03

R Kidney (g) M 0.19 ± 0.03 0.20 ± 0.02 0.15 ± 0.04 0.17 ± 0.04

F 0.20 ± 0.03 0.21 ± 0.03 0.15 ± 0.03 0.17 ± 0.03

Circ. = circumference, CRL = crown-to-rump length, g = grams, mm = millimetre, n = sample number, L =

left, R = right, P = postnatal day.

Appendix 5, Table 5. 1 Body weight, morphometry and organ weights at P14. Supplementary to Chapter 5

Section 5.4.1. Data of male and female pups are presented as Mean ± SD, and are analysed using a two-way

ANOVA with significance set at p < 0.05.

Appendix 5 (cont.)

Appendix 5, Table 5. 3 DEXA scan body composition measures at P14. Supplementary to Chapter 5,

Section 5.4.4. Data of male and female pups are presented as Mean ± SEM, and are analysed using a two-way

ANOVA with significance set at p < 0.05.

Control IUGR

Sex Saline n = 23 males

n = 24 females

DITPA n = 31 males

n = 18 females

Saline n = 33 males

n = 27 females

DITPA n = 34 males

n = 33 females

BMD (g/cm2) M 0.04 ± 0.001 0.04 ± 0.001 0.03 ± 0.001 0.04 ± 0.001

F 0.04 ± 0.001 0.04 ± 0.001 0.04 ± 0.001 0.04 ± 0.001

BMC (g) M 0.27 ± 0.023 0.27 ± 0.022 0.18 ± 0.019 0.19 ± 0.019

F 0.28 ± 0.024 0.31 ± 0.027 0.17 ± 0.022 0.19 ± 0.020

Bone Area (cm2) M 6.91 ± 0.48 6.73 ± 0.46 4.86 ± 0.40 5.03 ± 0.40

F 7.16 ± 0.49 2.82 ± 0.57 4.68 ± 0.46 4.99 ± 0.41

Lean tissue (g) M 22.57 ± 0.71 21.76 ± 0.68 18.29 ± 0.59 19.13 ± 0.59

F 22.16 ± 0.73 22.23 ± 0.85 17.74 ± 0.66 18.23 ± 0.61

Fat (g) M 9.56 ± 0.44 8.34 ± 0.42 6.94 ± 0.37 7.08 ± 0.37

F 9.79 ± 0.45 9.30 ± 0.53 7.26 ± 0.42 7.38 ± 0.38

% Fat M 29.87 ± 1.21 27.53 ± 1.15 27.39 ± 1.00 26.89 ± 1.00

F 30.74 ± 1.24 29.44 ± 1.45 28.44 ± 1.15 28.60 ± 1.04

BMD = bone mineral density, BMC = bone mineral content, g = grams, cm2 = centimetres squared, % =

percentage, n = sample number.

Control IUGR


n = 24 females

DITPA n = 31 males

n = 18 females

Saline n = 33 males

n = 27 females

DITPA n = 34 males

n = 33 females

Total brain (g) M 1.23 ± 0.07 1.24 ± 0.09 1.11 ± 0.08 1.11 ± 0.06

F 1.20 ± 0.06 1.18 ± 0.08 1.20 ± 0.07 1.09 ± 0.07

Brain: Body

(g/g)

M 0.03 ± 0.002 0.04 ± 0.005 0.04 ± 0.005 0.04 ± 0.004

F 0.03 ± 0.003 0.04 ± 0.005 0.04 ± 0.005 0.04 ± 0.008

Hemispheres M 0.91 ± 0.04 0.91 ± 0.06 0.80 ± 0.13 0.79 ± 0.16

F 0.89 ± 0.04 0.90 ± 0.06 0.79 ± 0.13 0.78 ± 0.08

Cerebellum M 0.16 ± 0.03 0.16 ± 0.03 0.14 ± 0.03 0.14 ± 0.02

F 0.15 ± 0.03 0.16 ± 0.03 0.14 ± 0.03 0.14 ± 0.03

Pons M 0.06 ± 0.01 0.06 ± 0.009 0.06 ± 0.01 0.05 ± 0.01

F 0.06 ± 0.008 0.06 ± 0.01 0.05 ± 0.01 0.05 ± 0.01

Medulla M 0.04 ± 0.002 0.04 ± 0.002 0.03 ± 0.002 0.03 ± 0.002

F 0.03 ± 0.002 0.04 ± 0.002 0.03 ± 0.002 0.03 ± 0.002

Appendix 5, Table 5. 2 Brain weights and brain-to-body weight ratio at P14. Supplementary to Chapter

5 Section 5.4.1 – 2.Data of male and female pups are presented as Mean ± SD, and are analysed using a two-

way ANOVA with significance set at p < 0.05.

Appendix 5 (cont.)

Appendix 5, Table 5. 4 Blood plasma assay results P14. Supplementary to Chapter 5, Section 5.5.6. Data

of male and female pups are presented as Mean ± SEM, and are analysed using a two-way ANOVA with

significance set at p < 0.05.

ALP = alkaline phosphatase, ALT = alanine transaminase, FT3 = plasma free triiodothyronine, FT4 = plasma

free thyroxine, n = sample number.

Control IUGR


n = 24 females

DITPA n = 31 males

n = 18 females

Saline n = 33 males

n = 27 females

DITPA n = 34 males

n = 33 females

FT3 M 1.95 ± 1.35 9.69 ± 1.20 2.41 ± 1.30 0.72 ± 1.21

F 2.41 ± 1.30 9.79 ± 1.51 0.72 ± 1.21 6.27 ± 1.21

FT4 M 25.18 ± 0.97 7.44 ± 0.90 24.92 ± 0.75 6.62 ± 0.73

F 31.21 ± 1.01 8.02 ± 1.01 25.76 ± 0.84 6.71 ± 0.79

ALT M 24.99 ± 3.30 30.01 ± 2.76 15.81 ± 3.02 24.29 ± 3.20

F 25.29 ± 3.85 26.24 ± 3.64 15.94 ± 2.97 22.53 ± 3.00

ALP M 344.56 ± 29.63 425.97 ± 24.80 386.02 ± 26.70 475.83 ± 27.74

F 315.75 ± 34.57 354.38 ± 32.57 354.80 ± 26.00 371.17 ± 26.24

Cholesterol M 4.39 ± 0.19 3.90 ± 0.16 4.05 ± 0.13 3.81 ± 0.15

F 4.24 ± 0.23 4.16 ± 0.23 4.05 ± 0.15 4.14 ± 0.14

Appendix 6

Monash Pathology Australia

FREE TRIIODOTHYRONINE (FT3) - Plasma Assay Protocol

The Access / DXI FreeT3 assay is a paramagnetic particle, chemiluminescent immunoassay for the quantitative

determination of free triiodothyronine levels in human plasma and plasma using the Beckman Coulter Unicel

DXI 800.

REAGENTS

Access Free T3 Reagent Pack Cat. No. A13422:

Access / DxI 800 Free T3 Calibrators Cat. No. A13430

ANALYTICAL RANGE

1.4 – 46 pmol/L

REFERENCE INTERVAL Plasma: 3.8 – 6.0 pmol/L

Source of Reference Interval

Beckman Coulter FT3 package insert

ANALYTICAL PERFORMANCE

BIORAD Immunoassay Plus levels 1, 2 and 3 (lyophilized) 4 x 5 mL of each level (Cat.No. 370)

MEAN SD CV % N

LEVEL 1 pmol/L 3.38 0.27 8.0 86

LEVEL 2 pmol/L 9.25 0.98 10.6 81

LEVEL 3 pmol/L 14.73 1.42 9.6 33

CLINICAL SIGNIFICANCE Triiodothyronine (3, 5, 3’-L- triiodothyronine, T3) is a hormone synthesized

and excreted from the thyroid gland, and formed by peripheral deiodination of thyroxine (T4). T3 and T4 are

secreted into the circulation in response to thyroid stimulating hormone (TSH) and play an important role in

regulating metabolism. The T3 and T4 secretion are regulated by a negative feedback mechanism involving

the thyroid gland, pituitary gland and hypothalamus. In the circulation 99.7% of T3 is reversibly bound to

transport proteins, primarily thyroxine binding globulin (TBG) and to a lesser extent albumin and prealbumin.

The remaining T3 does not bind to transport proteins but is free in circulation. This unbound fraction is

metabolically active. FT3 levels correlate with T3 secretion and metabolism. In hypothyroidism and

hyperthyroidism, FT3 levels parallel changes in total T3 levels. Measuring a FT3 level is useful when altered

levels of total T3 occur due to changes in T3 binding proteins especially TBG. TBG levels remain relatively

constant in healthy individuals but normal pregnancy and steroid therapy can alter these levels. In these

conditions the FT3 level is unchanged while the total T3 level parallels the changes in TBG.

REFERENCES i. Access Assay Manual (disc) Ref.387302 Version 3.0 ©2005 Beckman Coulter, Inc. Access Immunoassay Systems

FreeT3 Reagent Kit. REF. A13893B) package insert. Dated: 2005.


FREE THYROXINE (FT4) - Plasma Assay Protocol

PRINCIPLE / INTRODUCTION

The Access / DXI Free T4 assay is a two-step enzyme immunoassay carried out on a Beckman Coulter Unicel

DXI800. Monoclonal anti-thyroxine (T4) antibody coupled to biotin, sample, buffered protein solution, and

streptavidin-coated solid phase are added to the reaction vessel. During this first incubation the anti-T4

antibody coupled to biotin binds to the solid phase and the free T4 in the sample. After incubation in a reaction

vessel, separation in a magnetic field and washing removes any material not bound to the solid phase. Next,

buffered protein solution and triiodothyronine (T3)-alkaline phosphatase conjugate are added to the reaction

vessel. The T3-alkaline phosphatase conjugate binds to the vacant anti-T4 antibody binding sites. After

incubation in a reaction vessel, separation in a magnetic field and washing remove materials not bound to the

solid phase. A chemiluminescent substrate, Lumi-Phos* 530, is added to the reaction vessel and light generated

by the reaction is measured with a luminometer. The light production is inversely proportional to the

concentration of free T4 in the sample. The amount of analyte in the sample is determined from a stored, multi-

point calibration curve.

REAGENTS

• Access / DXI Free T4 Reagent Pack Cat. No. 33880

• Access / DXI Free T4 Calibrators Cat. No. 33885

ANALYTICAL RANGE

1.9 – 77.2 mol /L

Conversion factor: ng/dL x 12.9 = pmol/L

REFERENCE INTERVAL

0 – 4 days old 25 – 70 pmol/L

4 days - 6 months 12 – 30 pmol/L

>6 months 7.5 – 21.0 pmol/L

Source of Reference Interval

> 6 months; Beckman Coulter Kit Insert 386902B Dated 2005


The coefficient of Variation as assessed from routine quality control sera at three

Levels (Bio-Rad Ligand 1, 2, 3) is as follows:

MEAN SD CV % N

LEVEL 1 pmol/L 10.58 0.81 7.6 190

LEVEL 2 pmol/L 30.03 1.82 6.1 116

LEVEL 3 pmol/L 61.30 3.60 5.9 109

CLINICAL SIGNIFICANCE

Thyroxine (3, 5, 3’, 5’ tetraiodothyronine, T4) is a hormone synthesized and excreted from the thyroid gland.

T3 and T4 are secreted into the circulation in response to thyroid stimulating hormone (TSH) and play an

important role in regulating metabolism. The T3 and T4 secretion are regulated by a negative feedback

mechanism involving the thyroid gland, pituitary gland and hypothalamus. In the circulation 99.7% of T4 is

reversibly bound to transport proteins, primarily thyroxine binding globulin (TBG) and to a lesser extent

albumin and prealbumin. The remaining T4 does not bind to transport proteins but is free in circulation. This

unbound fraction is metabolically active. FT4 levels correlate with T4 secretion and metabolism. In

hypothyroidism and hyperthyroidism, FT4 levels parallel changes in total T4 levels. Measuring FT4 levels are

useful when altered levels of total T4 occur due to changes in T4 binding proteins especially TBG. TBG levels

remain relatively constant in healthy individuals but normal pregnancy and steroid therapy can alter these

levels. In these conditions FT4 levels are unchanged while total T4 levels parallel the changes in TBG.

REFERENCES

Access Assay Manual (disc) Ref.387302 Version 3.0 ©2005

Beckman Coulter Insert 386902B Access Free T4 Dated 2005


ALKALINE PHOSPHATASE (ALP) - Plasma Assay Protocol

PRINCIPLE / INTRODUCTION

The ALP method is an automated colorimetric method carried out on a Beckman Coulter SYNCHRON

LX20PRO System(s) with reagents and calibrators supplied by Beckman/Coulter. (Sydney, Australia) Cat

No 442670.

ALP reagent is used to measure alkaline phosphatase activity by a kinetic rate method using a 2-amino-2-

methyl-1-propanol (AMP) buffer. In the reaction, alkaline phosphatase catalyses the hydrolysis of the

colourless organic phosphate ester substrate, p-nitrophenylphosphate, to the yellow colored product, p-

nitrophenol, and phosphate. This reaction occurs at an alkaline pH of 10.3.

The SYNCHRON LX System(s) automatically proportions the appropriate sample and reagent volumes into

the cuvette. The ratio used is one part sample to 50 parts reagent. The system monitors the change in absorbance

at 410 nm. This change in absorbance is directly proportional to the activity of ALP in the sample and is used

by the System to calculate and express ALP activity.


The coefficient of variation as assessed from routine quality control sera at two levels.

BIORAD Liquichek Unassayed Chemistry Control (Human) Levels 1 and 2.

Cat. No 691 & 692.

MEAN SD CV% N

LEVEL 1 U/L 75.9 2.8 3.7 260

LEVEL 2 U/L 339.7 5.5 1.6 260

ANALYTICAL RANGE

5 - 1650 U/L

REFERENCE RANGES

30 - 120 U/L


Catalyses hydrolysis of orthophosphoric monoesters at pH 10. Has several isoenzymes: liver, bone and

placental. Increased in obstructive liver disease, osteoblastic or clastic bone disorders e.g. Metastatic,

hyperparathyroidism (primary or secondary). Decreased in hypophosphatasia.

REFERENCES

Beckman Coulter Chemistry Information Sheet 389874AA 4A ALP January 2021

Bowers, C.N. Jr. and McComb, R.B. Clin. Chem. 21: 1990 (1975).

N.W. Tietz, A.D. Rinker, L.M. Shaw: I.F.C.C. Method for Measurement of Catalytic Concentration of

Enzymes. Part 5. IFCC Method for Alkaline Phosphatase. J. Clin. Chem. Clin. Biochem. 21, no. 11 (1983).


ALANINE TRANSAMINASE (ALT) - Plasma Assay Protocol

Automated colorimetric method carried out on a Beckman Coulter SYNCHRON LX20PRO System(s)

with reagents and calibrators supplied by Beckman/Coulter. (Sydney, Australia) Cat No. 467840

The ALT- reagent is used to measure alanine transaminase in plasma or plasma by an enzymatic rate method.

In the assay reaction, the ALT catalyses the reversible transamination of L-alanine and alpha-ketoglutarate to

pyruvate and L-glutamine. The pyruvate is then reduced to lactate in the presence of lactate dehydrogenase

(LDH) with the concurrent oxidation of -Nicotinamide Adenine Dinucleotide (reduced form) (NADH) to -

Nicotinamide Adenine Dinucleotide (NAD).

The ALT- assay is based on the IFCC standard for enzyme determination. Pyridoxal-5'-phosphate is a cofactor

that is required for transaminase activity by binding to the enzyme using Schiff-base linkage.


a cuvette. The ratio used is one part sample to 11 parts reagent. The system monitors the rate of change in

absorbance at 340 nanometers over a fixed-time interval. This rate of change in absorbance is directly

proportional to the activity of ALT- in the sample and is used by the System to calculate and express the ALT-

activity.

One unit of enzyme activity is defined as the quantity of enzyme that catalyzes the reaction of 1 mol of

substrate per minute at +37C.

ANALYTICAL PERFORMANCE The coefficient of variation as assessed from routine quality control sera

at two levels.BIORAD Liquichek Unassayed Chemistry Control (Human) Levels 1 and 2.

Cat. No 691 & 692.

MEAN SD CV% N

LEVEL 1 U/L 24.63 2.14 8.69 144

LEVEL 2 U/L 86.69 2.73 3.14 155

ANALYTICAL RANGE

5 - 400 U/L

350 – 2600 U/L (ORDAC)

REFERENCE RANGE

< 3 years 5 - 45 U/L

Adult 7 - 56 U/L


ALT is a hepatic cytosolic enzyme which is more specific for this tissue than is AST. In liver disease associated

with hepatic necrosis e.g. viral hepatitis ALT is elevated before clinical signs and symptoms of disease e.g.

jaundice appear. Enzyme levels may reach 100 times the upper reference limit, but increases of 20 to 50 times

are more usual. Peak values are seen between 7 - 12 days, and levels return to normal by the third to fifth week

if recovery is uneventful. In toxic or viral hepatitis the ALT/AST ratio, which is normally less than 1,

approaches or becomes greater than 1. Moderate elevations of ALT are seen in extrahepatic cholestasis. Levels

of 5 to 10 times may be seen in primary or metastatic carcinoma of the liver. ALT may be slightly increased

after ingestion of alcohol, during delirium tremens, and after administration of drugs such as opiates,

salicylates, or ampicillin. Plasma elevations of ALT are rarely seen in conditions other than parenchymal liver

disease (although they may be occasionally increased in progressive muscular dystrophy and

dermatomyositis).

REFERENCES

Beckman Coulter Chemistry Information Sheet 389877AA 7A ALT-January 2021


CHOLESTEROL (CHOL) - Plasma Assay Protocol

The cholesterol was measured by a standard commercial enzymatic assay using a Beckman Coulter LX20PRO

Analyser, with reagents and calibrators supplied by Beckman Coulter Diagnostics Australia.

CHOL reagent is used to measure cholesterol concentration by a timed-endpoint method. In the reaction,

cholesterol esterase (CE) hydrolyses cholesterol esters to free cholesterol and fatty acids. Free cholesterol is

oxidized to cholestene-3-one and hydrogen peroxide by cholesterol oxidase (CO). Peroxidase catalyses the

reaction of hydrogen peroxide with 4-aminoantipyrine (4-AAP) and phenol to produce a colored quinoneimine

product.


the cuvette. The ratio used is one part sample to 100 parts reagent. The system monitors the change in

absorbance at 520 nanometers. This change in absorbance is directly proportional to the concentration of

CHOL in the sample and is used by the System to calculate and express CHOL concentration.

Cat No. 467825 Beckman Coulter Cholesterol Kit

CALIBRATOR: SYNCHRON Systems Lipid Calibrator

QUAILTY CONTROL: BIORAD Liquichek Unassayed Chemistry Control

(Human) Level 1 and Level Cat. No. 691 & 692

IMPRECISION: CV 1.9% @ 3.4 mmol/L

CV 1.3% @ 7.0 mmol/L

ANALYTICAL RANGE: 0.13 - 19.43 mmol/L

15.54 – 29.9 mmol/L (ORDAC)

REFERENCE RANGE: 0 - 5.5 mmol/L (NHF recommendation)

REFERENCES: Beckman Coulter Chemistry Information Sheet 38an Coulter Chemistry

Information Sheet 389895AA 5A CHOL January 2021\

Appendix 7

22 March 2017

Dr Mary Tolcos School of Health and Biomedical Sciences RMIT University

Dear Mary, Research &

Innovation

GPO Box 2476V Melbourne VIC

3001 Australia

AEC 1702: Using thyroid hormone-based therapies to repair and protect the brain in the intrauterine growth restricted rat.

I am pleased to advise that this project has been approved by the RMIT University Animal Ethics Committee (AEC) for the period from 22 March 2017 until 22 March 2020. An approved version of the application is

attached.

Animals

Your application has been approved to use up to n=60 rats (female Wistar rat dams) and up to n=480 rats

(Wistar rat offspring) over the duration of the project.

The use of animals in scientific procedures is strictly regulated by the Australian code for the care and use of

animals for scientific purposes. The above project is conducted under a Scientific Procedures and Premises

License.

Responsibilities of investigators

1. Dr Mary Tolocs

2. Miss Delphi E. Kondos-Devcic

3. Miss Aminath Azhan

4. Dr Tania Romano

5. Mrs Madhavi Khore

6. Ms Courtney Gilchrist Responsibilities of investigators are described in the Australian code for the care and use of animals for scientific

purposes (section 2.4). Investigators have a ‘personal responsibility for all matters that relate to the wellbeing of

animals that they use, including their housing, husbandry and care. This responsibility extends throughout the

period of use approved by the AEC until provisions are made for the animal at the conclusion of their use’ (s.2.4.1).

Amendments and extensions

If you find reason to amend your research method you should advise the AEC and prepare a request for minor

amendment form. Please note that the AEC may only deal with ‘minor’ amendment requests. Major amendments to

projects normally require a new project application.

Adverse events or unexpected outcomes

As the primary investigator you have a significant responsibility to monitor the research and to take prompt steps to

deal with any unexpected outcomes. You must notify the AEC immediately of any serious or unexpected adverse

effects on animals, or unforeseen events, which may affect the ethical acceptability of your project.

Unwell animals must be immediately reported via the care forms available at the RMIT Animal Facility. In the case

of any emergency, the Animal Welfare Officer, may be contacted on 0409 521 234 at any time. In case of any

unexpected animal death, the researcher has a responsibility to organise an autopsy so as to determine the cause

of death.

Investigator guidelines for record keeping

Investigators are required to adhere to the strict guidelines regarding record keeping for their project. Note that

records associated with a project ‘should be available for audit by the institution and authorised external reviewers’.

Failure to maintain proper records may result in a compliance breach of the Code and place at risk the researcher’s

capacity to carry out research with animals.

Conditions of approval

The AEC may apply conditions of approval beyond the submission of annual/final reports. There are no specific

conditions attached to this project, except that described elsewhere in this letter.

Reports

Approval to continue a project is conditional on the submission of annual and final reports. Annual reports are

requested in December each year, and must be submitted whether or not the project has commenced or is inactive.

Report forms are available at www1.rmit.edu.au/staff/research/researchintegrity-and-governance/animal-ethics.

Failure to submit reports will mean that a project is no longer approved, and/or that approval will be withheld from

future projects.

All reports or communication regarding this project are to be forwarded to the research ethics coordinator at

[email protected]

On behalf of the AEC I wish you well with your research.

Dr Brad Hayward Research Ethics Coordinator On behalf of RMIT Animal Ethics Committee

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