TERATOGENIC AND ENDOCRINE- DISRUPTING EFFECTS OF...
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TERATOGENIC AND ENDOCRINE- DISRUPTING EFFECTS OF HYPOXIA ON DEVELOPMENT OF ZEBRAFISH (DANIO RERIO) SHANG HUI HUA DOCTOR OF PHILOSOPHY CITY UNIVERSITY OF HONG KONG NOVEMBER 2005
Transcript of TERATOGENIC AND ENDOCRINE- DISRUPTING EFFECTS OF...
TERATOGENIC AND ENDOCRINE-
DISRUPTING EFFECTS OF HYPOXIA
ON DEVELOPMENT OF ZEBRAFISH
(DANIO RERIO)
SHANG HUI HUA
DOCTOR OF PHILOSOPHY
CITY UNIVERSITY OF HONG KONG
NOVEMBER 2005
CITY UNIVERSITY OF HONG KONG
香港城市大學
Teratogenic and Endocrine-Disrupting Effects of Hypoxia on Development of Zebrafish
(Danio rerio) 缺氧對斑馬魚發育的致畸效應及內分泌干擾效應
Submitted to Department of Biology and Chemistry
生物及化學系 in Partial Fulfillment of the Requirements
for the Degree of Doctor of Philosophy 哲學博士學位
by SHANG HUIHUA
尚惠華
November, 2005 二零零五年十一月
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ABSTRACT
Hypoxia/anoxia caused by eutrophication and organic pollution occurs over
thousands of km2 and has caused mass mortality, population decline and major
changes in community structure and function in many aquatic ecosystems
worldwide. Hypoxia is now being considered as one of the major threats to aquatic
ecosystems, and the problem is expected to worsen in the coming years.
In this thesis, the zebrafish (Danio rerio) was employed as a model species to test the
hypotheses that hypoxia is teratogenic to fish embryos, and may affect sex
determination via endocrine disruption during fish development. Malformation,
growth, apoptosis pattern, gonad development, hormones and sex ratio were
measured, and temporal and spatial responses of hypoxia-inducible genes, genes
controlling the synthesis of sex hormones, as well as genes controlling apoptosis in
fish exposed to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1) were compared
over time to elucidate molecular mechanisms underpinning the effects observed.
Heart rate was initially increased when embryos were exposed to hypoxia, but was
followed by a rapid decrease after 96 hours post fertilization (hpf) (t-test, p<0.001).
It appears that enhancing oxygen uptake and maintaining oxygen delivery were
employed only as short-term strategies to deal with hypoxia, while reducing energy
expenditure was used as a long term strategy to better survive under hypoxic
conditions.
Embryos exposed to hypoxia showed a delay in their development, and body length
of fish in the hypoxic treatment was 12.3% shorter than their counterparts’ in the
normoxic control after 168 h of development (t-test, p<0.001). Upon exposure to
hypoxia, embryos lost synchronization in their development, with their tails
developing much faster than their heads. Skeletal deformities, predominantly
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manifested as altered axial spinal curvature, were clearly evident in the hypoxic fish.
After 168 h, percentage of malformation in the hypoxic treatment was significantly
higher (+77.4%) than that of the normoxic control (t-test, p<0.01). Concomitantly, a
significant reduction in percentage of apoptotic cells in the tails (-63.7%), and an
increase in percentage of apoptotic cells in the brain (+116%), were found in hypoxic
embryos, indicating that malformation may be mediated through the alteration of the
normal apoptotic pattern. In addition, a higher percentage (+121.1%) of embryos in
the hypoxic group failed to develop their vascular systems when compared with the
normoxic control, and died after 3 to 5 days.
Hypoxia retards fish growth and gonad development. After two months, body length,
body weight, gonad weight and gonadosomatic index (GSI) in female were
significantly reduced by hypoxia (t-test, p<0.01). After 4 months, body length, body
weight, gonad weight and GSI were all significantly reduced in both hypoxic
females and males. Histological examinations further confirmed that gonadal
development in both males and females was retarded after exposure to hypoxia for 4
months.
Hypoxia disrupts the balance of sex hormones at very early stages of fish
development. At 48 hpf, levels of testosterone (T) significantly increased while
estradiol (E2) concentrations significantly decreased in hypoxic fish, resulting in a
marked increase (+357%) of the testosterone/estradiol (T/E2) ratio in sexually
undifferentiated fish (t-test, p<0.05), but this pattern was reversed at 120 hpf. After 2
months of development, T and E2 were significantly reduced in hypoxic males (t-test,
p<0.01, p<0.001, respectively), but not in hypoxic females. After 4 months, an
increase in T was clearly observed in hypoxic females (t-test, p<0.05). The increased
T/E2 ratio observed in hypoxic females but not hypoxic males indicated that
disruption of sex hormones was more severe in hypoxic females than that in hypoxic
males, although sex differentiation and sexual development in both sexes were
similarly affected by hypoxia. The sex ratio of zebrafish was altered upon chronic
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exposure to hypoxia, resulting in a male-biased population in the F1 generation
(74.1±1.7% males in the hypoxic group vs 61.9±1.6% males in the normoxic groups;
Chi-square test, p<0.05). Vitellogenin (VTG) was significantly reduced by hypoxia
in both females (t-test, p<0.01 for 2 months and p<0.001 for 4 months) and males
(t-test, p<0.001 for 2 months and p<0.01 for 4 months) at 2 and 4 months, indicating
that oocyte development and egg production were also inhibited by hypoxia.
Expression of various sex hormone control genes (viz. VTG, 3β-HSD, CYP11A,
CYP19A and CYP19B) were determined at 10dpf, 40dpf, 2 months and 4 months.
At the onset of gonad differentiation at 10 dpf, all selected genes were significantly
down-regulated by hypoxia (t-test, p<0.05 for VTG and p<0.001 for other genes). At
40 dpf, when sex differentiation and sex reversal occurred, VTG was up-regulated
(t-test, p<0.001) while all other genes were down-regulated (t-test, p<0.05).
Expression of CYP11A, CYP19A, CYP19B, VTG, and 3β-HSD were altered at
different developmental stages and in both sexes, showing that hypoxia can disrupt
key steps in sex hormone synthesis in both developing and adult fish. Our results
therefore indicate that synthesis of sex hormone and CYP aromatase in zebrafish
were disrupted by hypoxia during sexual differentiation and development, which
may account for the male-biased ratio observed in the hypoxic treatment.
HIF-1a was significantly up-regulated by hypoxia at 2 hpf (t-test, p<0.001), but
down-regulated thereafter. This cascaded into subsequent up-regulations of EPO,
VEGF (fold-changes ranging from 1.4±0.3 to 5.0±0.2 and 1.2±0.1 to 5.7±0.2,
respectively) and p53 at later stages of development. In particular, the ratio of
Bax/Bcl-2 in hypoxic fish was twice as high as that in normoxic fish at all time
points except 24 hpf, indicating that apoptosis was promoted in hypoxic fish. The
overall results suggested that 48hpf was the most sensitive window, at which time all
of the genes studied responded to hypoxia.
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At 36 hpf, up-regulation of Bax coupled with down-regulation of Bcl-2 was found in
the head region; while the reversed pattern was observed in the tail, suggesting a
higher apoptotic potential in the head region as compared to the tail region. This
offered further evidence to support the differential apoptotic patterns observed earlier
in hypoxic embryos. At two months, the ratio of Bax/Bcl-2 in hypoxic males was
198.2% of that measured in hypoxic females, implicating that apoptosis in males
might potentially be more susceptible to hypoxic effects than in females.
For the first time, this study provided experimental evidence to show that hypoxia is
a teratogen, which may induce premature death, growth retardation, malformation
and functional defects in zebrafish. We also report, for the first time, that hypoxia
can change the activity of P450 aromatase and the expression of various genes
controlling the synthesis of sex hormones, which in turn disrupt the balance of
testosterone and estradiol during fish development and sex differentiation, resulting
in a male-biased F1 generation. The increase in males and reduction of females in
fish populations caused by hypoxia may subsequently reduce reproductive success in
natural populations. Taken together with an increase in incidences of malformation
and pre-mature death, we conclude that hypoxia poses a significant threat to the
sustainability of natural fish populations over large areas worldwide. Because of the
similarity in genome and organ development between zebrafish and other mammals
including human and also because of the large scale and high frequency of
occurrence of hypoxia in the natural environment, it is possible that hypoxia may
also cause similar effects to other species.
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TABLE OF CONTENTS
Abstract……………….……………………………...………………………………..i
Declaration…………………………………………………………………………. vi
Acknowledgements………………………………………………………………… vii
Table of Contents…………………………………………………………………... ix
List of Figures……………………………………………………………………… xv
List of Tables……………………………………………………………………..... xxii
Acronyms and Abbreviations……………………………………………………..
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CHAPTER 1 INTRODUCTION………………………………………………….
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1.1 Hypoxia in the aquatic environment………………………………………. 1
1.2 Adaptive responses of aquatic organisms to hypoxia: behavioral,
physiological and biochemical responses………………………………….
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1.3 Molecular responses to hypoxia…………………………………………… 7
1.3.1 General molecular responses to hypoxia…………………………….
1.3.2 HIF-1α…………………………………………………………………
1.3.3 VEGF………………………………………………………………......
1.3.4 EPO…………………………………………………………………….
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1.4 Effects of hypoxia on embryonic development…………………………… 14
1.4.1 Effects of hypoxia on human embryonic development………..........
1.4.2 Effects of hypoxia on mammalian embryonic development…….....
1.4.3 Effects of hypoxia on chicken embryonic development…………….
1.4.4 Effects of hypoxia on amphibian embryonic development…...........
1.4.5 Effects of hypoxia on fish embryonic development…………………
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1.5 Effects of hypoxia on reproduction………………………………………... 19
1.5.1 Effects of hypoxia on human reproduction………………………….
1.5.2 Effects of hypoxia on mammalian reproduction ……………………
1.5.3 Effects of hypoxia on amphibian reproduction……………………...
1.5.4 Effects of hypoxia on fish reproduction………………………...........
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1.6 Effects of hypoxia on sex hormones……………………………………......
1.6.1 Effects of hypoxia on human sex hormones…………………………
1.6.2 Effects of hypoxia on mammalian sex hormones……………………
1.6.3 Effects of hypoxia on fish sex hormones…………………………......
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1.7 Apoptosis……………………………………………………………………. 25
1.7.1 Apoptosis……………………………………………………………….
1.7.2 Apoptosis and development………………………………………......
1.7.3 Hypoxia and apoptosis……………………………………………......
1.7.4 Apoptosis control genes……………………………………………….
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1.8 Sexual development………………………………………………………… 30
1.8.1 Sex steroid hormones in mammals…………………………………...
1.8.1.1 Testosterone…………………………………………………….
1.8.1.2 Estradiol……………………………………………………......
1.8.2 Sex steroid hormones in fish………………………………………….
1.8.3 Sex hormones and reproduction in fish……………………………...
1.8.4 Sex determination and differentiation in fish……………………….
1.8.5 Aromatase……………………………………………………………..
1.8.6 Sex hormone control genes……………………………………………
1.8.6.1 CYP11A (P450 scc) ……………………………………………
1.8.6.2 3β-hydroxysteroid dehydrogenase (3β-HSD)………………..
1.8.6.3 CYP19 (Aromatase)……………………………………….......
1.8.7 Vitellogenesis and vitellogenin (VTG)……………………………….
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1.9 Zebrafish……………………………………………………………………. 49
1.10 Research hypothesis and objectives……………………………………...... 51
CHAPTER 2 MATERIALS AND METHODS………………………………......
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2.1 Experimental fish…………………………………………………………… 53
2.1.1 Zebrafish as the study model………………………………………… 53
2.1.2 Zebrafish maintenance and embryo collection……………………... 54
2.2 Experimental design………………………………………………………... 55
2.3 Oxygen range finding experiment…………………………………………. 58
2.3.1 Viability……………………………………………………………….. 58
2.3.2 Retardation……………………………………………………………. 58
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2.4 Set-up of hypoxic and normoxic systems………………………………….. 59
2.5 Viability and biometry……………………………………………………… 61
2.5.1 Viability assay…………………………………………………………. 61
2.5.2 Biometry assay………………………………………………………... 61
2.5.2.1 Body length and growth of embryos and larvae…………...... 61
2.5.2.2 Body length, body weight and condition factor of adults…… 62
2.6 Effects of hypoxia on zebrafish development…………………………....... 62
2.6.1 Heart rate……………………………………………………………... 62
2.6.2 Malformation assessments…………………………………………… 63
2.6.3 Apoptotic pattern……………………………………………………... 63
2.6.3.1 Acridine Orange staining……………………………………... 63
2.6.3.2 Apoptotic pattern detection…………………………………… 64
2.7 Endocrine disrupting effects of hypoxia on zebrafish……………………. 66
2.7.1 Gonad weight and GSI……………………………………………...... 66
2.7.2 Hormone levels measurement……………………………………....... 66
2.7.2.1 ELISAs………………………………………………………….
2.7.2.2 Sample pre-treatment for sex hormone assay……………......
2.7.2.3 Testosterone…………………………………………………….
2.7.2.4 Estradiol………………………………………………………...
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2.7.3 Vitellogenin (VTG)……………………………………………………. 71
2.7.4 Gonad histology………………………………………………………. 72
2.7.5 Sex ratio……………………………………………………………...... 73
2.8 Expression of relevant genes under hypoxia……………………………… 74
2.8.1 Real-time RT-PCR…………………………………………………….
2.8.2 Sex hormone control genes and VTG………………………………...
2.8.2.1 Selection of genes and time points for detection…………......
2.8.2.2 Primers and probes…………………………………………….
2.8.3 Hypoxia inducible genes and apoptosis control genes………………
2.8.3.1 Selection of genes and time points for detection…………......
2.8.3.2 Primers and probes…………………………………………….
2.8.4 RNA extraction………………………………………………………..
2.8.5 DNAse I digestion of RNA……………………………………………
2.8.6 First-strand cDNA synthesis………………………………………….
2.8.7 Real-time RT-PCR reagents and cycling…………………………….
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2.9 Statistical analysis…………………………………………………………...
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CHAPTER 3 RESULTS……………………………………………………………
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3.1 Oxygen range finding experiment…………………………………………. 86
3.1.1 Viability………………………………………………………………...
3.1.2 Retardation…………………………………………………………….
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3.2 Viability and biometry……………………………………………………… 90
3.2.1 Viability assay………………………………………………………….
3.2.2 Biometry assay………………………………………………………...
3.2.2.1 Body length of embryos and larvae…………………………...
3.2.2.2 Body length, body weight and condition factor of adults……
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3.3 Effects of hypoxia on zebrafish development……………………………... 94
3.3.1 Heart rate……………………………………………………………...
3.3.2 Malformation …………………………………………………………
3.3.3 Apoptotic pattern……………………………………………………...
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3.4 Endocrine disrupting effects of hypoxia on zebrafish……………………. 99
3.4.1 Gonad weight and GSI……………………………………………......
3.4.2 Sex hormones………………………………………………………….
3.4.2.1 Sex hormone levels in embryos and larvae ………………….
3.4.2.1.1 Testosterone………………………………………......
3.4.2.1.2 Estradiol………………………………………………
3.4.2.2 Sex hormone levels in juveniles and adults………………......
3.4.2.2.1 Testosterone………………………………………......
3.4.2.2.2 Estradiol………………………………………………
3.4.2.3 Ratio of testosterone/estradiol………………………………...
3.4.3 Vitellogenin (VTG)…………………………………………………….
3.4.4 Gonad histology……………………………………………………….
3.4.4.1 Testis…………………………………………………………….
3.4.4.2 Ovary……………………………………………………………
3.4.5 Sex ratio……………………………………………………………......
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3.5 Expression of relevant genes under hypoxia……………………………… 117
3.5.1 Sex hormone control genes and VTG…………………………….......
3.5.2 Hypoxia inducible genes and apoptosis control genes………………
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3.5.2.1
3.5.2.2
3.5.2.3
3.5.2.4
3.5.2.5
Temporal change of expression patterns………………......
Spatial change of gene expression pattern in head and tail
Change of gene expression pattern between males and
females……………………………………………………….
Bax/Bcl2 ratio………………………………………………..
Correlation between expression patterns of various genes.
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CHAPTER 4 DISCUSSION…………………………………………………….....
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4.1 Mortality and biometry…………………………………………………….. 140
4.2 Effects of hypoxia on zebrafish development…………………………....... 141
4.2.1 Malformation………………………………………………………….
4.2.2 Heart rate and vascular formation……………………………….......
4.2.3 Apoptosis……………………………………………………………....
4.2.4 Summary……………………………………………………………....
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4.3 Endocrine disrupting effects of hypoxia on zebrafish…………………..... 148
4.3.1 GSI…………………………………………………………………......
4.3.2 Histological study………………………………………………….......
4.3.3 Sex ratio……………………………………………………………......
4.3.4 Endocrine disruption during embryonic development…………......
4.3.5 Endocrine disruption during sexual development…………………..
4.3.5.1 Sex hormone levels……………………………………………..
4.3.5.1.1 Testosterone…………………………………………..
4.3.5.1.2 Estradiol………………………………………………
4.3.5.2 Sex hormone balance and aromatase…………………………
4.3.6 Vitellogenin (VTG)………………………………………………….....
4.3.7 Summary……………………………………………………………....
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4.4 Expression of relevant genes under hypoxia…………………………….... 158
4.4.1 Sex hormone control genes and VTG…………………………….......
4.4.2 Hypoxia inducible genes and apoptosis control genes………………
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4.4.2.1 HIF-1α…………………………………………………………..
4.4.2.2 VEGF…………………………………………………………...
4.4.2.3 EPO……………………………………………………………..
4.4.2.4 P53…………………………………………………………........
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4.4.2.5 Bax, Bcl-2 and Bax/Bcl-2……………………………………… 168
4.4.3
4.4.4
4.4.5
Correlation between different hypoxia inducible genes and
apoptosis control genes…………………………………………......
Sensitivity to hypoxia in different developmental stages…………
Summary………………………………………………………….....
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4.5 Sex-dependent effects of hypoxia…………………………………….......... 173
4.6 Hypoxic biomarkers………………………………………........................... 176
4.6.1 Traditional biomarkers……………………………………….............
4.6.1.1 Malformation………………………………………..................
4.6.1.2 Sex ratio………………………………………...........................
4.6.1.3 GSI………………………………………...................................
4.6.1.4 VTG………………………………………..................................
4.6.2 Potential biomarkers……………………………………….................
4.6.2.1 Bax/Bcl-2………………………………………..........................
4.6.2.2 Testosterone/estradiol……………………………………….....
4.6.3 Potential biomarkers for hypoxia………………………………….....
4.6.3.1 HIF-1α………………………………………..............................
4.6.3.2 VEGF………………………………………...............................
4.6.3.3 EPO………………………………………..................................
4.6.3.4 Apoptotic pattern………………………………………............
4.6.4 Summary………………………………………....................................
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4.7 Ecological implications………………………………................................... 185
4.8 Overall conclusion………………………………………………….............. 188
BIBLIOGRAPHY………………………………………………………................. 190
APPENDIX………………………………………………........................................ 247
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LIST OF FIGURES
Fig. 1.1 Effects of anthropogenic activities on oxygen levels in aquatic systems…………………………………………………………………
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Fig. 1.2 General adaptive strategies to aquatic hypoxia (summarized and modified from Wu, 2002)………………………………………….......
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Fig. 1.3 Principal molecular responses to hypoxia and related gene regulation ………………………………………………………….......
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Fig. 1.4 Structure of testosterone (T)……………………………………….....
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Fig. 1.5 Structure of estradiol (E2)…………………………………………......
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Fig. 1.6 Hypothalamus-pituitary-gonad (HPG) axis and main reproductive hormones in fish. GnRH: gonadotropin- releasing hormone; GtH: gonadotropins (including FSH and LH); FSH: follicle- stimulating hormone; LH: luteinizing hormone; E2: estradiol; T: testosterone; 17 α , 20β DP: 17α, 20β-dihydroxy-4-pregnen-3-one; 11-KT: 11-ketotestosterone; VTG: vitellogenin (modified from Kime, 1999)……………………………………….…………………………...
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Fig. 1.7 Sex determination and differentiation in mammals (after Greenstein, 1994)………………………………………………………
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Fig. 1.8 Steroidogenesis in male fish……………………………………….......
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Fig. 1.9 Steroidogenesis in female fish…………………………………………
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Fig. 1.10 Hormonal control of vitellogenin (VTG) synthesis in female fish (Sumpter & Jobling, 1995) …………………………………………...
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Fig. 1.11 Development of zebrafish…………………………………………......
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Fig. 1.12 Hypotheses postulated in the present project……………..................
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Fig. 2.1 Developmental stages of zebrafish……………………………………
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Fig. 2.2 Experimental design………………………………………………......
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Fig. 2.3 Set-up of the hypoxic control system…………………………………
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Fig. 2.4 Structure of Acridine Orange (acridinium chloride hemi- (zinc chloride)) ………………………………………………………………
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Fig. 2.5 Detection of apoptosis in zebrafish using acridine orange. (A), control. (B), treated with caffeine at 15 µg/ml. (arrow). (www.phylonix.com/apoptosis.html)....................................................
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Fig. 2.6 General ELISA procedures…………………………………………...
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Fig. 2.7 Procedure of quantitative real-time RT-PCR (Reverse Transcription - Polymerase Chain Reaction)……………………......
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Fig. 2.8 Protocol of PCR (Annealing temperature = 60°C)……………….....
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Fig. 3.1 (A) Cumulative mortality in zebrafish embryos caused by different oxygen levels, 5.8, 1.0, 0.8, and 0.5 mg O2 l-1, at 24, 48, 72, 96, 120, 168 hpf.. 150 eggs per tank, (N=5, Mean ± SE). Values that are significantly different from the control are indicated by asterisks (Student’s t-test: *, p<0.05, **, p<0.01, ***, p<0.001); (B) Percent mortality of 120 hpf zebrafish embryos under different oxygen levels……………………………………………………………………
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Fig. 3.2 Zebrafish embryos exposed to (A) normoxia (5.8 mg O2 l-1) and (B) hypoxia (0.5 mg O2 l-1), showing retardation of development in the latter………………………………………………………….………...
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Fig. 3.3 Number of surviving fish at 50 dpf and 120 dpf upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=5, Mean ± SE.) Values significantly different from the control are indicated by asterisks (Student’s t-test: **, p<0.01, ***, p<0.001)…………….
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Fig. 3.4 Body length of embryos/larvae at 48, 72, 96, 120 and 168 hpf upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=10, mean ± SE). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05, ***, p<0.001). …………………………………………………………….…
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Fig. 3.5 Body length and body weight of female and male zebrafish after (A) 60 days of development and (B) 120 days of development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=12, Mean ± SE). Values significantly different from the normoxic control are indicated by asterisks (t-test: **, p<0.01; ***,
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p<0.001). ……………………………………………………….………
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Fig. 3.6 Condition factor of female and male zebrafish after (A) 60 days and (B) 120 days of development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=12, Mean ± SE). No significant difference was found. (t-test, p>0.05).……...................….
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Fig. 3.7 Heart rate in: (A) zebrafish embryos and larvae at 48, 72, 96, 120, 168, 288 hpf upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=10, Mean ± SE); (B) normoxic control fish and malformed embryos/larvae from hypoxic group at 48, 72, 96 and 120 hpf, (N=6, Mean ± SE). Values significantly different from the control are indicated by asterisks (t-test: **, p<0.01; ***, p<0.001).……………………………………………....………………..
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Fig. 3.8 Typical examples of malformation caused by hypoxia (0.8 mg O2 l-1) at 48 hpf, 72 hpf and 96 hpf. ……………………………………...
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Fig. 3.9 Percentage of malformation in zebrafish embryos / larvae at 8, 72, 96, 120 and 168 hpf upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=5, Mean ± SE). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05; **, p<0.01)…………………………….……………
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Fig. 3.10 Mean numbers of apoptotic cells per fish identified by Acridine Orange staining at 24 hpf in zebrafish embryos upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=10, Mean ± SE). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05; **, p<0.01) ………………..
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Fig. 3.11 (A) Gonad weight and (B) GSI of female zebrafish after 60 days of development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=12, Mean ± SE). Values significantly different from the normoxic control are indicated by asterisks (t-test: **, p<0.01). …………………………………………………….
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Fig. 3.12 Gonad weights of (A) female and (B) male, and GSIs of (C) female and (D) male zebrafish after 120 days of development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=12, Mean ± SE). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05; ***, p<0.001). ……………………………………………………………….
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Fig. 3.13 Level of T and E2 (pg/ml) in zebrafish embryos (300 embryos for
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each replicate, pooled) at (A) 48 hpf and (B) 120 hpf upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SE). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05; **, p<0.01; ***, p<0.001). ……………………………………………...…
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Fig. 3.14 Ratio of T/E2 in zebrafish larvae at 40 dpf upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SE). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05)……………………………...
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Fig. 3.15 Ratio of T/E2 in female and male zebrafish at (A) 60 dpf and (B) 120 dpf of development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SE). Values significantly different from the normoxic control are indicated by asterisks (t-test: **, p<0.01; ***, p<0.001). ……………………….……………
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Fig. 3.16 VTG level in (A) female and (B) male zebrafish at 60 dpf and 120 dpf upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=5, Mean ± SE). Values significantly different from the normoxic control are indicated by asterisks (t-test: **, p<0.01; ***, p<0.001). ……………………………………………………………….
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Fig. 3.17 Gonad of normoxic (5.8 mg O2 l-1) male zebrafish after 120 days of development; SPG: spermatogonia; SPC: spermatocytes; SPD: spermatids; SPA: spermatozoa……………………………………….
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Fig. 3.18 Gonad of hypoxic (0.8 mg O2 l-1) male zebrafish after 120 days of development; SPG: spermatogonia; SPC: spermatocytes; SPA: spermatozoa……………………………………………………………
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Fig. 3.19 Percentage of spermatogonia (SPG), spermatocytes (SPC) and spermatids (SPD) in the testes of zebrafish after 120 days of development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=12-13, Mean ±SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: ***, p<0.001)…………………………………………………...
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Fig. 3.20 Percentage of oogonia (Oo) and previtellogenic (PreV), vitellogenic (Vit) and preovulatory oocytes (PreO) in female zebrafish after 120 days of development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=12-15, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: ***, p<0.001)………….……………………………..
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Fig. 3.21 %-male in zebrafish examined at 120 dpf and in all fish examined
at 60dpf plus 120 dpf upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=5, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (Chi-square test, *, p<0.05)……………………………………………
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Fig. 3.22 Expression of sex hormone control genes and VTG in zebrafish at (A) 10 dpf and (B) 40 dpf upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05; ***, p<0.001)…………………………...
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Fig. 3.23 Expression of sex hormone control genes and VTG of (A) female and (B) male zebrafish at 60 dpf of development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05; **, p<0.01; ***, p<0.001)…
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Fig. 3.24 Expression of sex hormone control genes and VTG of (A) female and (B) male zebrafish at 120 dpf of development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05; **, p<0.01; ***, p<0.001). ……………………………………………………....……….
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Fig. 3.25 Temporal change of HIF-1α expression in zebrafish upon exposure to hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). Asterisk(s) indicate values significantly different from the normoxic control (t-test: **, p<0.01; ***, p< 0.001); Letters indicate data from the same/different group (one way ANOVA, p<0.05) ……………...……
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Fig. 3.26 Temporal change of VEGF and EPO expression in zebrafish upon exposure to hypoxia (0.8 mg O2 l-1) (N=4, Mean ± SD). * indicates expression of VEGF significantly different from the normoxic control (t-test: **, p<0.01; ***, p< 0.001); # indicates expression of EPO significantly different from the normoxic control (t-test: #, p<0.05; ###, p< 0.001)…………………………….…………………….
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Fig. 3.27 Expression of HIF-1α, VEGF, EPO, p53, Bax and Bcl-2 in zebrafish at (A) 24 hpf and (B) 48 hpf upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: **, p<0.01; ***, p<0.001)…………….
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Fig. 3.28 Expression of HIF-1α, VEGF, EPO, p53, Bax and Bcl-2 in
zebrafish at 120 hpf during development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05; **, p<0.01; ***, p<0.001)…
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Fig. 3.29 Expression of HIF-1α, VEGF, EPO, p53, Bax and Bcl-2 in zebrafish at (A) 10 dpf and (B) 40dpf during development upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05; **, p<0.01; ***, p<0.001).…………………………………………………
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Fig. 3.30 Expression of HIF-1α, VEGF, EPO, p53, Bax and Bcl-2 in the (A) head and (B) tail of zebrafish upon exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1) at 36 hpf, (N=4, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: **, p<0.01; ***, p<0.001)…………….
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Fig. 3.31 Expression of HIF-1α, VEGF, EPO, p53, Bax and Bcl-2 in head and tail of zebrafish upon exposure to hypoxia (0.8 mg O2 l-1). N=4, Relative Mean ± SD after normalization to β-actin; * indicates values significantly different from the normoxic control (t-test: *, p<0.05; **, p<0.01; ***, p< 0.001); # indicates significant difference in expression of a specific gene between head and tail, as revealed by one way ANOVA (p<0.05)………………………………………….
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Fig. 3.32 Expression of HIF-1α, VEGF, EPO, p53, Bax and Bcl-2 in (A) females and (B) males after 60 days of exposure to normoxia (5.8 mg O2 l-1) and hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: *, p<0.05; **, p<0.01; ***, p<0.001)…………….….
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Fig. 3.33 Expression of HIF-1α, VEGF, EPO, p53, Bax and Bcl-2 in males and females after 60 days of exposure to hypoxia. N=4, relative Mean ± SD after normalization to β-actin; * indicates values significantly different from the normoxic control (t-test: *, p<0.05; **, p<0.01; ***, p<0.001); # indicates significant difference between males and females, as revealed by one way ANOVA (p<0.05) ………………………………………………………………..
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Fig. 3.34 Bax/Bcl-2 ratio in zebrafish at different time points during development upon exposure to normoxia (5.8 mg O2 l-1) and
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hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). Values significantly different from the normoxic control are indicated by asterisks (t-test: ***, p<0.001). Different letters indicate significant differences at different time points as revealed by one way ANOVA (p<0.05). ……………………………………………………………….
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Fig. 3.35 Bax/Bcl-2 ratio in head and tail regions of zebrafish at 36 hpf upon exposure to hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). A value significantly different from the normoxic control is indicated by asterisks (t-test: ***, p<0.001)……………………….………………..
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Fig. 3.36 Bax/Bcl-2 ratio in males and females after 60 days of development upon exposure to hypoxia (0.8 mg O2 l-1), (N=4, Mean ± SD). A value significantly different from the normoxic control is indicated by asterisks (t-test: **, p<0.01)………………………………………..
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Fig. 4.1 Changes in sex hormone control gene expression, ratio of T/E2 and CYParom with respect to key stages of gonad development in zebrafish exposed to hypoxia (0.8 mg O2 l-1) as compared with those developed under normoxia (5.8 mg O2 l-1) (A): at 10 dpf; (B) at 40 dpf; (C) at 60 dpf in male and female; (D) at 120dpf in male and female. (--): expression below detection limit; T/E2
a: ratio of testosterone/estradiol. Nb: no significant difference between normoxic control and hypoxic treatment……………….……………
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LIST OF TABLES
Table 1.1 Key mammalian steroid hormones and their primary functions..
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Table 2.1 Zebrafish embryo medium (pH 7.2) (Westerfield, 1995)…………
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Table 2.2 Danieau’s solution (pH=7.6) (Nasevicius & Ekker, 2000)………...
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Table 2.3 Purification protocol for ELISA assay…………………………….
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Table 2.4 The primer sequences for sex hormone control genes and vitellogenin…………………………………………………………...
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Table 2.5 The primer sequences for hypoxia inducible genes and apoptosis control genes……………………………………………....................
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Table 2.6 Protocol for DNAse I digestion of RNA preparation……………...
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Table 2.7 Protocol for first-strand cDNA synthesis…………………………..
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Table 3.1 Serum sex hormone levels (Testosterone and Estradiol) of zebrafish at 40 dpf, 60 dpf and 120 dpf upon exposure to normoxia (5.8 mg of O2 l-1) and hypoxia (0.8 mg of O2 l-1)………..
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Table 3.2 Correlation coefficients (r) between the various hypoxia inducible genes, apoptosis control genes and Bax/Bcl-2…………..
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Table 4.1 Differential effects of hypoxia on male and female zebrafish at 60 dpf as compared with the normoxic control of the respective sex…………………………………………………………………….
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Table 4.2 Differential effects of hypoxia on male and female zebrafish at 120 dpf as compared with the normoxic control of the respective sex…………………………………………………………………….
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ACRONYMS AND ABBREVIATIONS
11-KT 11-ketotestosterone
17α, 20β DP 17α, 20β-dihydroxy-4-pregnen-3-one
3β-HSD 3-hydroxysteroid dehydrogenase
AChE Acetylcholinesterase
AE Androstenedione
ANOVA Analysis of variance
AO Acridine orange, acridinium chloride hemi-(zinc
chloride)
ATP Adenosine triphosphate
CCD Charge-coupled device
cDNA Complementary deoxyribonucleic acid
CF / K-factor Condition factor
CNS Central nervous system
CV Coefficient of variation
d Day
DDT Dichloro-diphenyl-trichloroethane
DHEA Dehydroepiandrosterone
DHT Dihydrotestosterone
DNA Deoxyribonucleic acid
DO Dissolved oxygen
dpf Day post fertilization
E2 Estradiol
EE 17α-ethinylestradiol
ELISAs Enzyme-linked immunosorbent assays
EPO Erythropoietin
FSH / GtH I Follicle-stimulating hormone
GD Gestation day
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GH Growth hormone
GLUT Glucose transporter
GnRH Gonadotropin-releasing hormone
GSI Gonado-somatic index
GtH Gonadotropin
h Hour
H&E Hematoxylin and eosin
HIF-1 Hypoxia inducing factor 1
HIF-1α Hypoxia inducing factor 1α
HIF-1β Hypoxia inducing factor 1β
hpf Hour post fertilization
HPG Hypothalamus-pituitary-gonad
HSD Hydroxysteroid dehydrogenase
hTERT Human telomerase reverse transcriptase gene
ICM Intermediate cell mass
LH / GtH I Leutinizing hormone
mRNA Messenger ribonucleic acid
MT 17α-methyltestosterone
NP 4-nonylphenol
O2 Oxygen
Oo Oogania
PCR Polymerase chain reaction
PreO Preovulatory oocyte
PreV Previtellogenic
PRL Prolactin
RNA Ribonucleic Acid
RT-PCR Reverse transcription polymerase chain reaction
SD Standard deviation
SE Standard error
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SPA Spermatozoa
SPC Spermatocytes
SPD Spermatid
SPG Spermatogonia
T Testosterone
T3 Triiodothyronine
T4 Thyroxine
TMB Tetramethylbenzidine
TUNEL Terminal deoxynucleotide transferase- mediated
dUTP - digoxygenin nick-end-labeling staining
UV Ultraviolet
VEGF Vascular endothelial growth factor
VEGFR-1 / flt-1 Vascular endothelial growth factor receptor-1
VEGFR-2 Vascular endothelial growth factor receptor-2
Vit Vitellogenic
VTG Vitellogenin
yr Year
Zf-VTG Zebrafish vitellogenin
