MORPHOLOGICAL AND PHYSIOLOGICAL DEVELOPMENTAL CONSEQUENCES
OF PARENTAL EFFECTS IN THE CHICKEN EMBRYO (Gallus gallus domesticus)
AND THE ZEBRAFISH LARVA (Danio rerio)
Dao H. Ho, B.A., M.S.
Dissertation Prepared for the Degree of
DOCTOR OF PHILOSOPHY
UNIVERSITY OF NORTH TEXAS
August 2008
APPROVED:
Warren Burggren, Major Professor and Dean
of the College of Arts and Sciences
Thomas Beitinger, Committee Member
Edward Dzialowski, Committee Member
Pamela Padilla, Committee Member
Paul Sotherland, Committee Member
Art Goven, Chair of the Department of
Biological Sciences
Sandra L. Terrell, Dean of the Robert B.
Toulouse School of Graduate Studies
Ho, Dao H. Morphological and physiological developmental consequences of
parental effects in the chicken embryo (Gallus gallus domesticus) and the zebrafish
larva (Danio rerio). Doctor of Philosophy (Biology), August 2008, 166 pp., 4 tables, 31
illustrations, references, 227 titles.
Cardiac, metabolic and growth response of early-stage chicken embryos to
perturbations in yolk environment was investigated. Also, effects of parental hypoxia
exposure on hypoxia resistance, thermal tolerance and body length of zebrafish larvae
were investigated.
In the first study, thyroxine, triiodothyronine and testosterone produced
differential effects on heart rate and development rate of chicken embryos during the
first 4 days of development. Triiodothyronine caused a dose-dependent increase in
heart rate when applied at 40 or 70 hours of age, while thyroxine caused a dosedependent increase in heart rate when applied at 40 hours only. Testosterone and
propyl-thiouracil (deiodinase antagonist) did not have an effect on heart rate.
Development rate was not changed by thyroxine, triiodothyronine, testosterone or
propyl-thiouracil, which suggested that heart rate changes did not result from changes
in embryo maturity. In the second study, chicken embryos exposed to yolks of different
bird species during early-stage embryonic development showed changes in heart rate,
mass-specific oxygen consumption and body mass that scaled with the egg mass,
incubation period length, and yolk triiodothyronine and testosterone levels of the
species from which yolk was derived. In the third study, this phenomenon was
investigated between layer and broiler chickens. Heart rate, oxygen consumption and
body mass of broiler and layer embryos were significantly changed by a breed-
specific change in yolk environment. Yolk triiodothyronine and testosterone
concentrations of broiler and layer eggs did not suggest that these hormones were
responsible for physiological and morphological changes observed. The final study
demonstrated that hypoxia resistance and body lengths, but not thermal tolerance of
zebrafish larvae was increased by parental hypoxia exposure.
Coypyright 2008
by
Dao H. Ho
ii
ACKNOWLEDGEMENTS
I would like to thank my committee members, Thomas Beitinger, Ed Dzialowski,
Pam Padilla and Paul Sotherland for their contributions and guidance during my
graduate career here at the University of North Texas. I would like to thank Susan
Chapman for her assistance with the development of the culture method used in my
experiments. Also, I would like to thank Wendy Reed and Todd Boonstra for their
collaboration and work on the radioimmunoassays, and their kind assistance and
hospitality during my stay in Fargo, ND. Thanks to Angie Trevino, Miho Fisher, Sheva
Khorrami, Jessie Brown, Adam Cochrane, Bonnie Myer, Francis Pan, Greta Bolin, Tara
Blank, Matt Gore, Kelly Reyna, Hunter Valls and Lindsey Taylor for their friendship and
support throughout the five years of long hours in the lab. Thanks to Dan Ball for his
whole-hearted support and kindness. Special thanks to my family: Mom, Dad, Amy,
Doris, Linh and Tuan who have put up with me for the last twenty-nine years, and
especially the last five years. Last, but definitely not least, I would like to thank Warren
Burggren for his instrumental role in advancing my growth and development as a
researcher – look, I‟m spinning plates!
The research was supported by NSF award number: 0128043, and SICB GrantIn-Aid of Research award.
iii
TABLE OF CONTENTS
Page
ACKNOWLEDGMENTS…………………………………………………………………..…..iii
LIST OF TABLES…………………………………………………………………………..…..v
LIST OF FIGURES……………………………………………………………………………..vi
Chapter
1. INTRODUCTION……………………………………………………………..…….1
2. CHRONOTROPIC CARDIAC EFFECTS OF TRIIODO-L-THRYONINE,
L-THYROXINE, PROPYL-THIOURACIL AND TESTOSTERONE IN
EARLY-STAGE CHICKEN EMBRYOS………………..………………….…….18
3. INFLUENCE OF INTER-SPECIES YOLK FACTORS ON HEART RATE,
OXYGEN CONSUMPTION AND BODY MASS OF EARLY-STAGE CHICKEN
EMBRYOS………………..…….…..……………………...………………………48
4. INFLUENCE OF INTER-BREED YOLK FACTORS ON HEART RATE,
OXYGEN CONSUMPTION, DEVELOPMENT RATE AND BODY MASS OF
EARLY-STAGE CHICKEN EMBRYOS………………..………….……………86
5. INFLUENCE OF PARENTAL HYPOXIA EXPOSURE ON STRESS
SURVIVAL PHYSIOLOGY OF ZEBRAFISH LARVAE….…………………...116
6. PARENTAL EFFECTS DURING DEVELOPMENT OF VERTEBRATES:
CONCLUSIONS AND FUTURE DIRECTIONS………………………….……143
REFERENCES…………………………………………………………….…………………148
iv
LIST OF TABLES
3.1 Egg components and length of prenatal development (incubation period) of
birds.....................................................................................................................74
3.2 Least-squares regression estimation on log10-transformed data for the relationship
between yolk hormone concentrations (yolk T3 or yolk TE) and IEM………….…76
3.3 Least-squares regression estimation on log10-transformed data for the relationship
between IP and yolk hormone concentrations (yolk T 3 or yolk TE)…...………….76
3.4 Morphological and physiological traits of White Leghorn chicken (Gallus gallus)
cultured on yolks of different bird species……………………………………….….79
v
LIST OF FIGURES
2.1 Mean relative heart rate of chicken embryos exposed to T 4 (0, 2.5, 5, 10 or 20
ng/embryo) at 40 h, or at 70 h of age………………………………………………..37
2.2 Mean relative heart rate of chicken embryos exposed to T 3 (0, 2.5, 5 and 10
ng/embryo) at 40 h or at 70 h of age…………………………………………...……38
2.3 Mean relative heart rate of chicken embryos exposed to T (0, 2.5, 5, 10 or 20
ng/embryo) at 40 h or at 70 h of age………………………………………..……….39
2.4 Mean percent change in heart rate over time of chicken embryos exposed to T 4
(0, 2.5, 5, 10 or 20 ng/embryo) at 40 h of age……..……………………………….40
2.5 Mean percent change in heart rate over time of chicken embryos exposed to T 3
(0, 2.5, 5 or 10 ng/embryo) at 40 h of age……………………………………....….41
2.6 Mean percent change in heart rate over time of chicken embryos exposed to
PTU (0, 15, 30 or 60 ug/embryo) at 40 h of age……………..…………………….42
2.7 Mean percent change in heart rate over time of chicken embryos exposed to
TE (0, 2.5, 5, 10 and 20 ng/embryo) at 40 h of age………………….……………43
2.8 Mean stage of chicken embryos exposed to T 4 (0, 2.5, 5, 10 or 20 ng/embryo)
at 40 h or at 70 h of age…………………………..…………………………………..44
2.9 Mean stage of chicken embryos exposed to T 3 (0, 2.5, 5, 10 or 20 ng/embryo)
at 40 h or at 70 h of age…………………………………………..…………………..45
2.10 Mean stage of chicken embryos exposed to PTU (0, 15, 30 or 60 µg / embryo)
at 40 h or at 70 h of age……………………………...……………………………….46
2.11 Mean stage of chicken embryos exposed to TE (0, 2.5, 5, 10 or 20 ng/embryo)
at 40 h or at 70 h of age…………………………………………………………..…..47
3.1 Schematic of embryo culture method……………………………………..……………73
3.2 Relationship between initial egg mass (IEM) and incubation period (IP) for 7
species of birds…………………………………………………………….…………..75
3.3 Relationship between log initial egg mass (IEM) and log yolk T 3 or log yolk TE
for 7 species of birds……………………………………………………….………….77
3.4 Relationship between log yolk T3 or log yolk TE and log IP for 7 species
of birds……………………………………………………………………………….....78
vi
3.5 Relationship between log body mass (BM) and log yolk T3, log yolk TE,
log IEM, or log IP for chicken embryos cultured on the yolks of 7 species
of birds…………………………………………………………………………………..80
3.6 Relationship between log heart rate (fH) and log yolk T3, log yolk TE,
log IEM, or log IP for chicken embryos cultured on the yolks of 7
species of birds………………………………………………………………………...82
3.7 Relationship between log mass-specific oxygen consumption rate ( ) and
log yolk T3, log yolk TE, log IEM, or log IP for chicken embryos cultured
on the yolks of 7 species of birds…………………………………………………….84
4.1 Wet and dry body mass of broiler and layer embryos cultured on broiler
or layer egg yolk for a period of 12, 24 and 48 hours after the start of
experiments………………………………………………………………………..….110
4.2 Developmental stage broiler and layer embryos cultured on broiler or layer egg
yolk for a period of 12, 24 and 48 h after the start of experiments…..…..……..111
4.3 Mass-specific oxygen consumption of all cultured embryos, and broiler and
layer embryos cultured on broiler or layer egg yolk as a function of body
mass…………………………………………………………………………………...112
4.4 Mean heart rate of broiler and layer embryos cultured on broiler or layer egg yolk
for a period of 12, 24 and 48 hours………………………………………...………113
4.5 Yolk T3 and TE concentrations of broiler and layer eggs……………………..…….114
4.6 Egg components mass of broiler and layer eggs as a function of egg mass….…115
5.1 Hypoxia resistance of larval offspring of zebrafish exposed to hypoxia or
normoxia for 1, 2, 3 or 4 weeks………………….………………………………….136
5.2 Hypoxia resistance of larval offspring of zebrafish exposed to hypoxia or
normoxia for 1, 2, 3 or 4 weeks among clutches………………….……….……..137
5.3 Critical thermal limits of zebrafish larvae of parents exposed to normoxia or
hypoxia for 12 weeks………………………………………………………...………138
5.4 Mean body length of larval offspring of zebrafish exposed to hypoxia or
normoxia for 1, 2, 3 or 4 weeks……………………………………………………..139
5.5 Mean body length of larval offspring of zebrafish exposed to hypoxia or
normoxia for 1, 2, 3 or 4 weeks among clutches…………………………..……..140
vii
5.6 Fecundity of adult zebrafish exposed to normoxia or hypoxia for 1, 2, 6 and 12
weeks…………………………………………………………………….……………141
5.7 Egg and egg component volumes of eggs of adult zebrafish exposed to
normoxia or hypoxia for 1, 2, 6 or 12 weeks………………………………..……..142
viii
CHAPTER 1
INTRODUCTION
Research Objectives
I investigated epigenetic effects of maternally-derived yolk factors and parental
environment on the physiology and morphology of developing animals using two
vertebrates models: chicken and zebrafish. The chicken model was used primarily to
gain an understanding of the comparative nature of maternal effects of egg yolk factors
on early cardiovascular development (Chapters 2, 3 and 4).
The zebrafish model
research investigated the effects of parental hypoxia exposure on offspring stress
resistance (Chapter 5).
In the first study (Chapter 2), I examine heart rate and development rate of earlystage
chicken
embryos
exposed
to
thyroid
hormones
(thyroxine
(T 4)
and
triiodothryonine (T3)) and testosterone (TE) to determine how changes in hormone
levels can influence development during the first four days of life. I hypothesized that
hormones, such as T3, T4 and testosterone, which are present in yolks of all bird eggs,
can significantly alter heart rate of chicken embryos in a dose- and age-dependent
fashion.
In the second study (Chapter 3), I describe heart rate, mass-specific oxygen
consumption (
), and body mass of chicken embryos that were placed on the yolks of
different bird species, i.e., quail, turkey, duck, goose, ostrich and emu, during the initial
days of embryonic development to determine if early embryonic phenotype of chicken
embryos would be affected by a species-specific change yolk environment.
1
I also
describe egg mass, length of incubation period and yolk T3 and yolk TE concentrations
of the eggs of these various birds to explore whether changes in these factors covary
with the changes in the phenotype of chicken embryos developing on the yolks of
different species. I hypothesized that the inter-species variation in egg mass, length of
incubation period and yolk hormone concentrations would influence the physiological
and morphological phenotype of chicken embryos exposed to the yolks of different bird
species.
In the third study (Chapter 4), I assessed heart rate, mass-specific
,
development rate and body mass of broiler and layer chicken embryos that were placed
on different yolks (either broiler or layer egg yolks) during the first few days of
embryonic development to determine if early embryonic phenotypes of these two
chicken breeds would be affected by a breed-specific change in yolk environment. I
hypothesized that the exchange of yolks between broiler and layer embryos during early
development would cause the embryos to develop phenotypic traits that are
characteristic of the breed from which the yolk was derived.
In the fourth study (Chapter 5), I describe hypoxia resistance, thermal tolerance
and body length of zebrafish larvae born to parents exposed to hypoxic or normoxic
environments for variable lengths of time, to elucidate whether parental environment
significantly influences the physiology and morphology of the larval offspring. I also
describe the egg clutch size and egg component volumes in the adult zebrafish
exposed to hypoxia and normoxia to determine the possible mechanism of parental
effects.
I hypothesized that exposing parents to hypoxia would increase hypoxia
resistance in the zebrafish larvae, but not the thermal tolerance of these larvae.
2
Epigenetic Processes
Historically, the term “epigenesis” was coined to describe how phenotypes arise
from the genotype during the development of an individual (Waddington, 1957). As
understanding of genetics and hertiability advanced, the definition of epigenetics
underwent many revisions, and today remains disputed (Ptashne, 2007; Bird 2007).
Although controversy surrounds the issue of defining a “true” epigenetic effect, evidence
of epigenetic effects in physiology are widespread. Generally, epigenetics is the study
of biological consequences of gene function modification without concomitant change in
gene sequence. Mechanisms of epigenetic processes include, but are not limited to,
histone modifications, heterochromatin regulation, and DNA methylation (Holliday,
2006). The study of epigenetics has gained much attention in the past decade largely
due to the discovery that, in many instances, epigenetic processes outweigh direct
genetic processes in the manifestation of many aberrant phenotypes (Baylin and
Herman, 2000; Jones and Takai, 2001; Egger et al., 2004; Weinhold, 2006).
Parental effects, defined as the impact of the parental genome on the phenotype
of the offspring, is one of the most extensively studied epigenetic processes in
physiology (Fox and Mousseau, 1998).
Parental effects can occur during pre-
fertilization, fertilization and post-fertilization. Pre-fertilization parental effects principally
include imprinting.
Imprinting is the changing of offspring genetic expression via
methylation and/or histone modification of the offspring‟s chromatin by the parental
genome.
Post-fertilization effects involve an array of parentally-derived factors
conserved throughout the plant and animal kingdom. Parental effects have been shown
to be responsible for cellular organization, body axis determination, and general
3
performance of the organism prior to start of zygotic transcription, and in later stages of
development are influential in the initiation and maturation of organ systems (Rossiter,
1996; Fitch et al., 1998; Reinhold, 2002). Moreover, parental effects are long-lived, and
are detected even after the individual is removed from parental influence (Lacey, 1996;
Bernardo, 1996; Mousseau and Fox, 1998; Sinervo and Huey, 1999). Thus, parental
effects play a crucial role in the evolutionary process.
Maternal Effects in Oviparious Species
Parental effects can be further divided into two categories: maternal effects and
paternal effects. Of the two categories, maternal effects have been more intensely
investigated, perhaps due to the intimate role of the mother during offspring embryonic
and juvenile development in most animals. In oviparous animals, maternal effects that
occur after fertilization are the result of the protective barrier of an egg, and/or
maternally produced nutrient stores, mRNAs, transcription factors, immune factors,
antioxidants, and hormones deposited into the egg during oogenesis (Smith and Ecker,
1965; Craig and Piatigorsky, 1971; Rose and Orlans, 1981; Schwabl, 1993; Mousseau
and Fox, 1998; Kudo, 2000). The first studied maternal effect was in the pulmonate
snail, Limnaea peregra, where directionality of shell coiling was determined to be
inherited in a non-mendelian fashion via a yet-to-be discovered factor in the yolk
(Boycott and Diver, 1923; Sturtevant, 1923). Since then, the role of maternal effects in
early development has been studied in a wide variety of species (Bernardo, 1996; Yan,
1998; Eising et al., 2006). Synthesis of these studies strongly suggests that maternal
effects during early development can have a long-lasting impact on adult phenotype by
4
changing developmental trajectories, as well as having acute impact on embryo
function.
Although the impact of maternal effects in pre-zygotic tissue organization and
post-zygotic development has gone in and out of fashion throughout the past century, it
is an undeniable factor in evolution (Nüsslein-Volhardt et al., 1987; Bernardo, 1996;
Mousseau and Fox, 1998; Wolf et al., 1998; Wilson et al., 2005).
Of evolutionary
importance is the impact of maternal effects on life history traits of the offspring that
have a clear and direct bearing on fitness.
Examples range from age at first
reproduction, number and size of offspring to reproductive lifespan and aging. Ever
since Willham (1972) revised the heritability model to account for maternal effects,
researchers have demonstrated that maternal effects represent a significantly large
portion of the variation of traits such as offspring birth weight, offspring birth date, natal
litter size, and juvenile mass in mammalian species (McAdam et al., 2002; Wilson et al.,
2005). Maternal effects also account for variation in performance, stress resistance,
and life history traits between and within species in cases where the embryo heavily
relies on large maternally-produced yolk and albumen reserve for growth and
development (Bernardo, 1996; Sinervo and Huey, 1999; Finkler et al., 1998;
Pakkasmaa and Jones, 2002; Blanc et al., 2003; Dzialowski and Sotherland, 2004).
This suggests that, maternal effects not only are prevalent in oviparous species, but that
maternal effects can play a significant role in the evolutionary process.
Maternal investment during egg formation often includes creation of a protective
outer membrane/shell covering, and deposition of egg factors into the yolk and/or
albumen. Egg factor quality is wholly determined by phylogeny and maternal condition
5
(Davis and Ackerman, 1985).
Egg shell characteristics principally affect offspring
development by regulating the ionic and molecular composition of the environment
surrounding the embryo, and protecting the embryo within from biotic and abiotic
challenges (Romanoff, 1960).
In most cases, the outer covering of eggs is semi-
permeable, with the permeability of the membrane setting the influx rate of lifesustaining molecules, i.e. oxygen and water, and the efflux rate of by-products of
metabolism, i.e. carbon dioxide. In avian species, this shell characteristic, also known
as shell conductance, determines oxygen tensions in the air-cell, and the rate of water
loss during incubation of eggs (Okuda and Tazawa, 1988; Balkan et al., 2006). Since
shell conductance affects the internal environment of the egg, slight changes in shell
conductance can result in significant changes in development (Wagner-Amos and
Seymour, 2003).
Egg factors such as hormones, immunoglobulins, antioxidants, mRNAs and
nutritive molecules in the yolk and albumen, largely influence the development of
embryos. The general quantity of yolk, the spatial distribution of yolk factors, and the
overall concentration of yolk factors play an important role in the ultimate size, shape,
mass, performance and behavior of the individual (Rose and Orlans, 1981; Sinervo and
Huey, 1990; Schwabl, 1993; Mousseau and Fox, 1998; Kudo, 2000; Eising et al., 2006;
Saino et al., 2007).
The pattern of maternal mRNA distribution in the egg of D.
melanogaster largely determine body pattern and appendage placement (Winslow et
al., 1988), while the amount of yolk available to the developing embryo can influence
overall body growth and metabolism in birds and fishes (Heming and Buddington, 1988;
Dzialowski and Sotherland, 2004), and size and performance in lizards (Sinervo and
6
Huey, 1999). Furthermore, egg factor concentrations influence growth rate, juvenile
organ size and hatchling behavior in a variety of avian species (Scwabl, 1993;
Groothius and von Engelhardt, 2005; Schwabl et al., 2007).
The Avian Cardiovascular System
Development
The chicken heart is derived from cardiogenic cells located in the peripheral and
posterior portions of the blastoderm (the splanchic mesoderm); first appearing as two
lateral tube-like structures, then joining and becoming a single, ventro-medially located,
structure at the 7-8 somite stage (~30 hours incubation). During this stage the cardiac
tube begins to contract spontaneously (Olivo, 1924, 1925 in Romanoff, 1960).
Within
hours (11 somite stage; 38 hours), the tube beats as a unit, and becomes capable of
circulating blood. Blood and endothelial tissue formation in extra-embryonic tissues
occurs simultaneously with heart formation. Subsequently, blood islands fuse with the
embryonic vascular system, and form a continuous vascular network, the culmination of
which is known as the cardiovascular system (Romanoff, 1960).
Cardiac activity in early chicken embryos (3-4 days incubation) does not play a
role in overall growth or oxygen delivery; rather, it modulates/regulates cardiac
maturation and angiogenesis via hemodynamic (blood flow and pressure dynamics)
action (Burggren et al., 2000; Burggren, 2004). Thus, early cardiac function is crucial in
establishing the foundation of the cardiovascular system‟s subsequent role in
convective oxygen, nutrient, waste and hormone transport.
Blood transport in the
cardiovascular system depends upon blood pressure, peripheral resistance, and blood
viscosity. Each factor is dynamic and can be changed to modulate blood flow: 1)
7
pressure by cardiac output (determined by heart size, heart rate), 2) blood viscosity by
blood cell size and number, and 3) vessel parameters by angiogenic processes and
responses of endothelial walls. During development, these factors have the potential of
undergoing significant changes (both transient and permanent) in the face of epigenetic
processes (Schwerte et al., 2005). Dynamic changes in this system define the rate at
which delivery of life-sustaining factors occur, thus affecting the present and subsequent
growth and development of an organism.
Maternal Effects and the Cardiovascular System
The cardiovascular system is the first physiological system to become functional
in an organism, and thus, plays a key role in development of subsequent organ systems
(Burggren and Keller, 1997). The interplay of blood flow and pressure (hemodynamics)
plays a central role in angiogenesis, oxygen transport, and nutrient uptake.
Thus,
elucidating the factors that modulate cardiovascular development is crucial to
understanding and predicting the developmental trajectory of the whole organism. Of
particular interest here is how much embryonic genetic expression and maternallyderived factors contribute to early cardiovascular phenotypic variation among species.
Since Aristotle first observed the beating embryonic heart embryonic of the
chicken (Gallus gallus domesticus), the study of avian cardiovascular development has
revealed that the morphological and physiological variation that exist among individuals
of various avian species begins as early as the first few days of embryonic development
(Sellier et al., 2006). This suggests a direct genetic component in heart rate regulation,
but also allows the possibility of the role of indirect (maternal effect) genetic components
in early heart rate regulation (Burggren et al., 1994; Burggren, 1999; Burggren et al.,
8
2003; Schwerte et al., 2005). Although Reinhold (2002) reports that maternal effects in
birds are negligible, the traits included in the analysis were morphological and
behavioral in nature; cardiovascular physiology was not considered.
Avian species
progress through an extensive period of embryonic development within a maternally
produced egg which consists of a protective, semi-permeable shell, and a large supply
of yolk and albumen for growth and maturation. These maternal factors show marked
inter- and intra-specific differences in mass and yolk and albumen composition
(Groothuis et al., 2005). This strongly suggests a role for maternal effects in early
cardiovascular development of avian species. Thus, elucidating the role of maternal
effects in the developing cardiovascular system of birds seems prudent.
Comparative Embryonic Development in Birds
In addition to obvious phylogenetic differences in gross morphology among avian
species during late embryonic and hatchling stages, subtle physiological differences can
be observed during early embryonic development (Anthony et al., 1991). Embryonic
growth rate among species varies significantly, with incubation periods ranging from 13
to 56 days (Rahn et al., 1974). When examined inter-specifically, there is a strong
positive correlation between length incubation period and egg mass, and egg mass and
oxygen consumption, but there is a negative correlation between avian egg mass and
avian embryonic heart rate of various species at comparable stages of development
(Rahn et al., 1974; Tazawa et al., 1991). Some authors have suggested that shell
conductance is responsible for the negative inter-specific correlation between egg mass
and embryonic heart rate because shell conductance is negatively correlated with egg
mass (Rahn et al., 1974). However, inter-specific differences in cardiac performance
9
are also accompanied by inter-species differences in maternally-derived factors
deposited in yolk and albumen. Perhaps heart rate relationships among avian species
can be explained by these maternal factors.
Chickens are excellent models for studying the role of maternal factors in
variation of phenotypic traits because of the wide diversity of domestic chickens that
have been artificially selected for different purposes. To increase meat yield and egglaying efficiencies, the poultry industry has bred two types of chicken for these
purposes; the layer which is a prolific egg-layer and the broiler which has high meat
yield. Broilers are often four times heavier than layers at 42 days post-hatch, and have
significantly different muscle parameters (McRae et al., 2006). As a result of extremely
fast muscle tissue growth, broilers suffer from ascites, a syndrome that occurs when the
oxygen demands of muscle tissue exceeds the capabilities of the cardiovascular system
to deliver oxygen to these tissues (Olkowski, 2007).
Significant differences exist between the two chicken breeds not only in hatchling
and adult physiology and morphology, but also embryonic physiology and morphology
(Latimer and Brisbin, 1987; Burggren et al., 1994; Clum et al., 1995; Mitchell and Burke,
1995; Schreurs et al., 1995; Martinez-Lemus et al.., 1998; Martinez-Lemus et al., 1999;
Koenen et al., 2002; Odom et al., 2004). Embryonic mass differs significantly between
layers and broilers throughout incubation, with differences becoming more apparent
towards the end of incubation (Pal et al., 2002). Broiler embryos also have a faster rate
of development than layer embryos beginning at the first 40 hours of development
(Boerjan, 2005).
In fact, during the last 5 days of incubation, broiler embryonic
metabolic heat production becomes significantly higher than layer embryonic metabolic
10
heat production due to larger body size of broilers (Boerjan, 2005). However, when
body mass was taken into account, layer heat production per gram of body weight was
significantly higher than broiler heat production per gram of body weight (Sato et al.,
2006). Heart morphology also differs between the two breeds; with only broiler embryos
showing ventricular hypertrophy as early as 40 hours of incubation (Pas Reform, 1997).
Heart rate of broiler embryos are higher than that of layer embryos, but this difference is
reversed in post-hatch (Yoneta et al., 2006; Yoneta et al., 2007).
Undoubtedly, the morphological and physiological differences between broiler
embryos and layer embryos set the stage for the corresponding juvenile and adult
phenotypes. The question remains as to the contribution of maternally produced factors
in these phenomena. Although genetic differences do exist between the two breeds
(Dunnington, 1994; International Chicken Polymorphism Map Consortium, 2004), it is
not known what roles these genetic differences might play in determining the maternal
factors that influence the embryo during development. The first step in resolving this
issue is to empirically examine the apparent breed-specific differences in early
development between the broilers and layers in the context of breed-specific differences
in yolk factors.
Environmentally-Induced Parental Effects
The environment of an individual can have crucial bearing on not only its own
fitness, but also on that of its immediate offspring. This driving force of evolution is
termed, “environmentally-induced parental effect,” and is recognized as an epigenetic
process (Lacey, 1996). Environmental stress plays a role in adaptation and evolution
either by increasing rates of genetic mutation, modifying phenotypic variation, and/or
11
inducing the expression of novel phenotypes (Hoffman and Merilä, 1999; Badyaev,
2005). Under conditions of abiotic stress such as hypoxia, chemical, and radiation,
adults of a wide range of species show beneficial changes in parental investment and
care that promote advantageous offspring phenotype (Munkittrick and Dixon, 1988;
Meyer and Di Giulio, 2003; Mondor et al., 2005). Cotton aphids produced more winged
offspring when exposed to predator tracks, presumably to enhance the ability of their
offspring to escape predation (Mondor et al., 2005), while offspring of killifish and white
sucker fish of polluted environments produce offspring that are more resistant to the
specific pollutant (Munkittrick and Dixon, 1988; Meyer and Di Giulio, 2003). Along the
same lines, Henry and Harrison (2001) reported that in Drosophilidae melanogaster,
offspring of hypoxia-reared parents that were raised in normoxia expressed larger
tracheal systems, mimicking the phenotype of those individuals raised in hypoxia.
Larger tracheal systems may confer greater resistance to hypoxia, but this was not
directly tested. Moreover, the idea of “bet-hedging” by increasing variance in offspring
phenotype in the face of environmental stressors has gained support in the last decade.
In the plant, Arabidopsis thaliana, parental exposure to ultraviolet-C radiation caused an
increase in frequency of homologous recombination in the subsequent generation
(Molinier et al., 2006). The authors concluded that the epigenetic trans-generational
effect “may increase the potential for adaptive evolution.”
Hypoxic conditions are prevalent throughout environments of the past, present,
and the impending future; thus its role in acclimation, adaptation and evolution of
species has been, and is of great interest to the scientific community (Kasting, 2001;
Diaz, 2001). Currently, hypoxia affects a wide range of life forms. Terrestrial animals
12
often face low oxygen conditions due to low oxygen partial pressures at high altitudes,
and/or low oxygen concentrations in isolated atmospheric environments; while aquatic
animals are exposed to hypoxia due to a multitude of factors such as: 1) salinitydependent capacity of water for dissolved oxygen 2) temperature-dependent capacity
of water for dissolved oxygen and 3) oxygen-depleting, biotic activities such as low
levels of photosynthetic activity in bodies of water, and increased biotic load in
conjunction with decreased oxygen turn-over in ponds and lakes (Denny, 1990; Boyer
et al., 2006). Aquatic environments are increasingly at high risk of becoming hypoxic
due to human anthropogenic activities that contribute to the oxygen depleting factors
(Diaz, 2001), and the magnitude of anthropogenic activities show no signs of slowing
down.
Thus it is imperative to understand the impact of rapidly and drastically
decresing oxygen levels on the ability of aquatic animals and their offspring to survive.
Impact of Hypoxia on Organisms
Low oxygen availability induces transient, acute responses that often lead to
permanent changes within an animal‟s lifetime. Responses to hypoxia are somewhat
similar between mature and developing animals, but developing animals often display
unique and exaggerated responses due to the incomplete development of biological
systems (Burggren and Crossley, 2001).
These changes include morphological,
physiological, cellular, molecular and genetic alterations in traits that are paramount to
evolutionary fitness.
Adult Animals
Acute exposure to hypoxia often leads to immediate physiological responses
such as increased ventilation and increased blood flow via increased cardiac output and
13
vasodilation to maintain adequate oxygen supply to the tissue (Holeton and Randall,
1967; Burggren and Randall, 1978; Burleson et al., 2002).
If hypoxic stress is
prolonged, the resulting phenotypic changes include, but are not limited to, increased
hematocrit and hemoglobin levels that increase the oxygen carrying capacity of the
blood, increased surface area of respiratory structures (i.e. gills in fish, tracheal system
in insects, and skin in amphibians and larval fishes) and increased blood flow to
increase diffusion and capture of oxygen from the environment, and metabolic
depression to conserve energy (Feder, 1983a,b; Boutilier et al., 1988; West and Vliet,
1992; Dalla Via et al., 1994; Severi et al.., 1997; Sollid et al., 2003; Harrison et al.,
2006).
These adaptations not only increase the potential of hypoxia survival in the
individual during the time of hypoxia exposure, but also incur changes that can result in
increased survival during subsequent hypoxic stress as well as resistance to other
stressors.
For example, zebrafish (Danio rerio) pretreated to sub-lethal levels of
hypoxia had increased survival times in subsequent lethal hypoxia tests when
compared to non-pretreated fish (Rees et al., 2001).
Furthermore, in some cases
resistance (or exposure) to one stress confers resistance to a wide range of other
stresses. Thermal acclimation in adult C. elegans confers tolerance/resistance against
not only thermal stress, but hypoxia and heavy metal stress (Katschinksi and Glueck,
2003). The protein profile in organisms that are exposed to acute and chronic hypoxia
show many similarities to the profiles of organisms that are exposed to extreme
temperature and high levels of heavy metals (Airaksinen et al., 1998; Sørensen et al.,
14
2001; Deane and Woo, 2006). This strongly suggests a common underlying genetic
basis for stress resistance to a variety of abiotic factors.
As energy and resources are expended to mount resistance against hypoxia,
biological systems not directly involved in the survival of the individual tend to suffer.
Fish exposed to hypoxia often have reduced fitness due to low fecundity, decreased
mating rates, and low survivorship of offspring. Wu et al. (2003) reported that adult carp
exposed to moderate levels of hypoxia suffered disruption of the endocrine systems that
resulted in lowered hatchability of the offspring as well as lethal mutation phenotypes in
offspring.
Developing Animals
In
developing
animals,
hypoxia
exposure
induces
morphological
and
physiological changes similar to those observed in adult animals; but the caveat of a
developing organism is that incomplete development of biological systems can, and will
result in unique responses to abiotic stressors (Burggren and Crossley, 2001).
Although acute responses of developing animals to hypoxia include cardioventilatory
changes and haematological changes as seen in adult animals (Jacob et al., 2002; Jonz
and Nurse, 2005), hypoxia exposure during “sensitive” periods (critical windows) in
development often results in permanent changes in the juvenile and adult morphology
and physiology. Hypoxia exposure throughout development resulted in an increased
ratio of phenotypic females to phenotypic males in the affected population (Shang et al.,
2006), and also induces endocrine disruption in developing zebrafish, similar to the
disruption seen in adults (Wu et al., 2003; Shang and Wu, 2004). In D. melanogaster,
cell size and cell number of the wings and epithelium irreversibly decrease when the
15
individual is exposed to hypoxia up to the pupa stage (Peck and Maddrell, 2005; Farzin
et al., 2007).
Additionally, hypoxia exposure during development resulted in
preferentially larger tracheal size in the D. melanogaster (Henry and Harrison, 2004).
Although the relationship between size and hypoxia resistance/tolerance has not been
clarified, size has clear implications in oxygen demand/uptake (Feder, 1983; Burleson et
al., 2001; Robb and Abrahams, 2003; Ospina and Mora, 2004).
In addition to inducing morphological changes in developing animals, hypoxia
can alter the onset of the regulation of a number biological systems. Development of
brine shrimp (Artemia salina) respiratory regulation is accelerated by hypoxia, and as a
consequence, these animals suffer a decrease in Darwinian fitness (Spicer and ElGamal, 1999). Hatching occurs earlier in chicken embryos incubated under hypoxia
when compared to those incubated in normoxia (Blacker et al., 2004). Hypoxia also
delays the onset of cardiovascular regulation in zebrafish (Bagatto, 2005). The genetic
and molecular mechanisms underlying these physiological responses to hypoxia
challenge is correlated to Insulin-like Growth Factor (IGF) Binding Protein-1 expression
mediated by the hypoxia-inducible factor-1 (HIF-1; Maures and Duan, 2002; Kajimura et
al., 2006).
Alterations in gene expression involving these factors occur in both
developing and adult aquatic animals (Gracey et al., 2001; Ton et al., 2003; Nikinmaa
and Rees, 2005).
Hypoxia Survival in Zebrafish
Hypoxia resistance is an animal‟s ability to combat the lethal/damaging effects of
low oxygen levels by maintaining normal internal oxygen levels despite low ambient
levels, whereas hypoxia tolerance is an animal‟s ability to survive despite low internal
16
oxygen levels (Hochachka et al., 1996; Haddad, 2006).
During development from the
single-cell stage to the adult stage, the zebrafish progresses through a number of
modes of oxygen capture, thus requiring them to contend with hypoxia stress via very
different teleological “strategies.” Interestingly, two-cell stage zebrafish embryos are
able to tolerate anoxic conditions by entering into a state of suspended animation, with
this ability gradually disappearing by the time of hatching (Padilla and Roth, 2001). This
shows that tolerance to oxygen depletion is short-lived, and that hypoxia resistance (as
opposed to tolerance) is the key to zebrafish survival during the latter part of its
developmental period.
Prior to 12 days post-fertilization (dpf), oxygen supply to the
tissues occurs via diffusion across the body surface. At 2.5 thru 3 dpf, gill filament buds
begin to form on the pharyngeal arches, increasing the surface area of the fish (Kimmel
et al., 1995). Moreover, before 5 dpf, circulating red blood cells are suggested to serve
functions other than oxygen transport (Pelster and Burggren, 1996). Thus, during this
early period of development, a reasonable strategy for hypoxia-resistance would be to
increase the body surface area to volume ratio and/or increase the oxygen permeability
of the skin/body surface membrane, and/or increase water flow across diffusive
surfaces (Liem, 1981). As the larva increases in size and becomes constrained by
oxygen diffusion alone, the cardiovascular system begins to take on properties of
convective circulation (~14 dpf), and the gills become functional (Kimmel et al., 1995;
Jacob et al., 2002). It is during this time that cardioventilatory mechanisms combat the
effects of hypoxia (Jonz and Nurse, 2005; Jonz and Nurse, 2006; Vulesevic and Perry,
2006; Vulesevic et al., 2006).
17
CHAPTER 2
CHRONOTROPIC CARDIAC EFFECTS OF TRIIODO-L-THYRONINE,
L-THYROXINE, PROPYL-THIOURACIL AND TESTOSTERONE IN
EARLY-STAGE CHICKEN EMBRYOS
Introduction
In oviparous species, maternally-derived factors, such as hormones, are
deposited in physiologically-relevant amounts into the egg yolk and albumen during egg
formation, and thus have the strong potential to modulate directly the development of
offspring throughout embryogenesis (Groothuis and Schwabl, 2008). Altering levels of
maternally-derived hormones during embryogenesis can elicit not only immediate
morphological and physiological responses of embryos, but also lead to permanent
changes in juvenile and adult phenotype of the offspring (Rose and Orlans, 1981;
Heming and Buddington, 1988; Sinervo and Huey, 1990; Schwabl, 1993; Mousseau
and Fox, 1998; Kudo, 2000; Eising et al., 2006; Saino et al., 2007; Groothuis and von
Engelhardt, 2005; Schwabl et al., 2007; Groothuis and Scwabl, 2008). Moreover, yolk
hormone profiles show physiologically-relevant variations within and among oviparous
species. Thus, effects of maternally-derived egg factors on offspring phenotype have
been acknowledged to be a major driving force in evolutionary processes (McNabb,
2006; Garamszegi et al., 2007; McGlothlin and Ketterson, 2008; Martin and Schwabl,
2008). Previous studies of effects of maternally-derived
(and exogenously
administered) hormones on avian phenotype predominantly examined perturbations in
morphology during early embryonic development and changes in physiology and
behavior during late stages of development and adulthood. Rarely have physiological
18
effects of maternally-derived hormones been investigated at the time of early-stage
organogenesis, which is a vulnerable period of development in which dramatic
maturation and integration of physiological systems occurs in the context of heavy
maternal influence (McNabb and Wilson, 1997; Wingfield et al., 2008).
Little is known about the influence of yolk hormones on the developing
cardiovascular system. Early cardiac function is crucial in producing the foundation for
the subsequent role of the cardiovascular system in convective oxygen, nutrient, waste
and hormone transport (Burggren and Keller, 1997), and thusly deserves investigation
during early stages of development. The cardiovascular system of the chicken (Gallus
gallus domesticus) embryo undergoes drastic morphological and physiological
maturation over a short period of time (40 to 96 hours of incubation, Hamburger &
Hamilton stages 11 to 20 (HH); Hamburger and Hamilton, 1951), during which, the
embryonic endocrine system is not yet fully functional (McNabb and Wilson, 1997).
Presence of hormone receptors prior to the functioning of corresponding hormonal
systems in embryos strongly suggests the importance of maternally-derived hormones
during embryogenesis (McNabb and Wilson, 1997). Furthermore, thyroid hormones
(triiodothyronine (T3) and thyroxine (T4)) and steroid hormones, i.e. testosterone (TE))
exist at physiologically significant levels in the yolk of unincubated avian eggs, and have
been intensely studied due to their significant effects on offspring morphology,
physiology and behavior (McNabb and Wilson, 1997; Groothuis and Schwabl, 2008).
Thyroid hormones have direct chronotropic effects and cardioregulatory effects
involving cardiac cell proliferation and growth in developing and adult animals.
Hypothyroidism in adult rates resulted in smaller hearts and slower heart rates
19
(Balkman et al., 1992), whereas experimenter-induced hyperthyroidism in neonatal rats
caused significant hyperplasia and slight hypertrophy of heart ventricles (Slotkin et al.,
1992). Administration of T3 caused gross malformations in early chicken embryos (2-3
days of incubation), but caused no cardiac-specific morphological deformities (Flamant
and Samarut, 1998), while exogenous T4 administration at 3 days of incubation
increased hyperplasic and hypertrophic cardiac growth in 5 to 9 day old chicken
embryos (Schjeide et al., 1989). Treatment with propylthiouracil (PTU), a deiodinase
inhibitor, reduced the symptoms of hyperthyroidism by blocking conversion of the less
potent form of T4 to the more potent form of T 3 (Robbins, 1981), and reduced cardiac
cell proliferation and growth in developing systems (Moussavi et al., 1985). Additionally,
TE, a hormone primarily investigated for its role in maturation of sexual traits,
aggression, and overall juvenile body size, has recently been shown to promote
differentiation of embryonic stem cells into beating cardiomyocytes (Goldman-Johnson
et al., 2008). Despite strong evidence of the morphological and functional effects of T 3,
T4, PTU and TE upon the developing cardiovascular system, chronotropic cardiac
effects of these hormones on the early developing embryo remains unknown.
The
hormonally-dependent
the
maturation
and
relatively
early
development
of
cardiovascular system makes it an ideal system for investigating how maternallyderived hormones, per se, may influence physiological features of early embryos and,
possibly, the developmental trajectory of subsequent late-stage embryonic and juvenile
development.
I hypothesized that hormones (T 3, T4, and TE) that exist in physiologicallyrelevant levels in the avian egg environment can significantly affect the cardiac
20
development of the early developing embryo of Gallus gallus domesticus.
Also, I
hypothesized that the disruption in deiodinase function via PTU will reduce heart rate of
the early developing embryos. I used heart rate to reveal perturbations in early cardiac
development and function associated with alterations in T3, T4 and TE levels. As a
means of explaining whether the chronotropic effects of the aforementioned hormones
are attributable to cardiac-specific mechanisms, or to general development-promoting
mechanisms early in development, I also characterized the developmental rate of
embryos treated with these pharmacological agents. I assessed the morphological and
physiological variables in embryos dosed at two developmental periods to determine if
chronotropic effects were age-dependent this early in development.
Materials and Methods
Egg Storage and Incubation
Fresh White Leghorn chicken eggs were purchased from Texas A&M University
(College Station, TX). Upon arrival, eggs were stored at room temperature (23-26°C)
for no longer than 3 days before the start of incubation.
Eggs were incubated a
commercial incubator (NBS CO2 incubator, Model CO-21, New Brunswick Scientific,
Edison, NJ) held at 37.5 + 0.3°C and 58-68% humidity. Eggs were positioned blunt end
up, and eggs were not turned during the course of the incubation period (~33 hours (h))
to position embryos just underneath the air-cell at the time of windowing (see windowing
method below). Once windowed, eggs remained in the incubator for assessment of
heart rate (fH) and developmental rate.
21
Windowing of Eggs
Windowing consisted of cutting away the eggshell that overlies the air-cell (blunt
end), and removing the inner shell membrane to reveal the developing embryo. The
final window size was approximately the diameter of the air-cell of each egg (~3-4 cm).
The bottom of a 35 mm x 15 mm plastic Petri-dish was placed over the window to
provide a clear view of the embryo while reducing water loss. Windowed eggs were
then placed in a commercial Styrofoam incubator with a clear plastic top (Electronic
Hova-bator incubator, Model 1588, G.Q.F Manufacturing Co., Savannah, GA), held at
37.5 + 0.3 °C with 58-68% humidity for the remainder of the study.
Experimental Design
Both chronic and acute effects of T 3, T4, PTU and TE on embryonic heart rate
and developmental rate were determined. Chronic exposure experiments included an
added factor of age at the time of hormone exposure, which allowed for characterization
of hormone effects at two distinct ages, 64 h and 94 h. Chronic exposure experiments
consisted of administering the hormone to 40 h embryos or 70 h embryos, and
recording heart rate and stage of development 24 h post administration (64h and 94 h,
respectively).
Acute exposure experiments involved assessment of heart rate and
developmental stage for each embryo 1 h prior to administering of the hormone at 70 h
(baseline heart rates), and assessing heart rate and stage 1, 2, 3 and 6 h post
administration (hpa). For each pharmacological agent under investigation, at least two
trials were conducted for chronic and acute exposure experiments.
22
Administration of Pharmacological Agents
Drug Solutions
L-thyroxine (T4) sodium salt pentahydrate (Sigma T2752) was solubilized in 1N
sodium hydroxide (NaOH; 1 mg/0.05 ml), before diluting with 50 ml 1X Tyrode‟s
Solution to obtain a solution of 20 ng/µl. This solution was then serially diluted to obtain
the desired concentrations of 2.5, 5 and 10 ng/µl.
Triiodo-L-thyronine (T3) solution was made in the same fashion as T 4 solution,
except 0.5 mg of T3 (3,3‟, 5-triiodo-L-thryonine, Sigma P3755) was solubilized in 0.05 ml
1NaOH prior to mixing with 50ml 1X Tyrode‟s. This produced T3 concentrations of 2.5,
5 and 10 ng/µl. For both T3 and T4 solutions, an appropriate volume of 1N NaOH was
added to each dilution to maintain a concentration of 0.1% 1N NaOH.
Testosterone (Sigma T1500) was first solubilized in 70% ethanol (1 mg/0.5 ml)
prior to dilution with 50 ml 1X Tyrode‟s to obtain a solution of 20 ng TE/µl. This solution
was then serially diluted to obtain the desired concentrations of 2.5, 5 and 10 ng/µl. At
each dilution, an appropriate volume of 70% ethanol was added to each solution to
maintain a concentration of 0.7% ethanol.
Propyl-thiouracil was made by solubilizing 12mg PTU (6-propyl-2-thiouracil,
Sigma P3755) into 10ml of Tyrode‟s solution to obtain a concentration of 60 ug/50 ul.
This solution was then serially diluted to obtain additional solutions of 30 and 15 ug/50
ul.
Vehicle solutions consisted of either Tyrode‟s at 0.1% 1N NaOH (0 ng T 3/µl and
0 ng T4/ul), Tyrode‟s at 0.7% ethanol (0 ng TE/ul), or Tyrode‟s solution (0 ug PTU/50 µl).
All solutions were prepared 2 h prior to administration.
23
Doses of T3, T4 and TE chosen for this study reflect both physiological and
pharmacological levels of these hormones in avian yolks to ensure the complete
characterization of the chronotropic effects of the hormones (Schwabl, 1993; McNabb
and Wilson, 1997; Flamant and Samarut, 1998). PTU doses were determined based on
doses used in mid-stage chicken embryo studies since no PTU dose ranges were
available for early-stage embryos (Bargman and Gardner, 1967; Raheja and Linscheer,
1979).
Administration
Drug solutions were applied to windowed embryos, at either 40 (HH10- HH11) or
70 h of age (HH17-HH19) by topical application of 1 µl (T 3, T4 and T experiments), or 50
µl (PTU experiment) onto the embryo. This method of administration has been used in
numerous studies and found to be effective (Schjeide,1989; Flamant and Samarut,
1998; Henry and Burke, 1999; Daisley et al., 2005).
Heart Rate
Heart beat of embryos was visually counted over the course of 30 seconds (sec).
For each embryo, heart rate was an average of three recordings. If the heart rate
recordings for an embryo varied more than 50 bpm, the embryo was excluded from
statistical analyses. Heart rates were reported as beats per min (bpm). For embryos
dosed at 40 h of age, heart rates were measured at 24 hpa. For embryos dosed at 72 h
of age, heart rates were measured at 0, 1, 2, 3, 6, and 24 hpa.
Development Rate
In general, embryonic development rate was determined by staging embryos,
according to HH stages, at the start of the experiment and then again each time heart
24
rate measurements were taken. Assigning stage to embryos was primarily based on
the degree of curvature of the spine (HH14 and older). For example, stage HH14 was
indicated by the forebrain and hindbrain forming a 45° angle while HH15 was indicated
by 67.5° curvature of the forebrain and hindbrain.
Statistical Analyses
In chronic exposure experiments, heart rate for each trial was first converted to
change in heart rate from mean heart rate of the vehicle-treated group, relative to mean
heart rate of the vehicle group (f ∆H = ((fH –
(fH vehicle))/
(fH vehicle)),
and then arcsin
transformed for statistical analyses. Embryo stage data were considered to be discrete
and were ranked prior to analyses, but were converted back for presentation. Factorial
two-way analyses of variance (ANOVA) with respect to dose and administration age (40
h old and 70 h of age) were performed on f∆H and embryo stage for each
pharmacological agent.
Tukey‟s post-hoc analyses were used to compare values
among doses within each administration age, and between administration age at each
dose. Statistical significance was assigned a value of α < 0.05. Data were expressed
as means + S.E.M.
In acute exposure experiments, heart rate was first converted to the change in
heart rate from baseline heart rate relative to baseline heart rate (f ∆H = (fH
baseline)/
fH
baseline),
post admin.
– fH
and then arcsin transformed for statistical analyses. Baseline heart
rate is defined as the heart rate of each individual embryo 1 h prior to administration of
treatment. Separate factorial two-way ANOVAs with respect to dose and time elapsed
after administration (1, 2, 3 and 6 h) were performed on f ∆H for each drug experiment.
Tukey‟s post-hoc analyses were used to compare values among doses for each drug.
25
Statistical significance was assigned a value of α < 0.05. Data were expressed as
means + S.E.M. For both chronic and acute exposure experiments, heart rate data
were transformed back and converted to percentage for graphical presentation.
Results
Heart Rate
Baseline Heart Rates
Windowed chicken embryos incubated at 37.5°C and treated with vehicle
solution at volumes of 1 or 50 μl displayed mean heart rate of 189 + 5 bpm at 64 h of
age, and mean heart rate of 243 + 5 bpm for 94 h of age.
Chronic hormone exposure
Thyroxine.
Age significantly affected (F(1,
225)
= 23.09, p < 0.001) heart rate
response of embryos exposed to T4. Embryos dosed at 40 h showed a significant dosedependent increase in mean relative heart rate, whereas embryos dosed at 70 h
showed minimal chronotropic responses, in comparison to the vehicle-treated controls
(Fig. 2.1). Twenty-four after the administration of 10 and 20 ng/embryo at 40 h of age,
the mean relative heart rate of embryos were significantly elevated above vehicletreated embryos (Tukey‟s, q = 6.30, p < 0.001; q = 5.283, p = 0.002, respectively).
However lower doses of 2.5 and 5 ng/embryo did not induce any significant
chronotropic effects beyond vehicle controls. Although the mean relative heart rates of
embryos dosed with 10 and 20 ng T4/embryo at 70 h was not significantly different from
vehicle treated controls, they were significantly different from the mean relative heart
rate of embryos dosed at 40 h of age (Tukey‟s, q = 6.12, p < 0.001; q = 5.45, p < 0.001,
respectively).
26
Triiodo-L-thyronine. Administration of T3 at either 40 h or 70 h of age produced
similar dose-dependent significant increases in mean relative heart rate that, however,
did not differ from one another. Statistical significance was reached at a dose of 5
ng/embryo for embryos dosed at 70 h old (Tukey‟s, q = 4.54, p = 0.01; Fig. 2.2). Effects
of higher doses (10 ng/embryo) were not significantly different from vehicle-treated
controls.
Propyl-thiouracil. PTU administration produced no significant changes in mean
relative heart rate at any dose (0, 15, 30 or 60 ug / embryo) within any administration
age (40 and 70 h old.)
Testosterone.
Administration of TE to 40 h or 70 h embryos produced no
chronotropic effects when compared to the respective vehicle groups (Fig. 2.3).
However, when compared between administration ages, the mean relative heart rate of
embryos administered 2.5 and 5 ng/embryo at 40 h of age were significantly higher than
those of embryos administered the same doses at 72 h (Tukey‟s, q = 3.59, p = 0.013; q
= 2.93, p = 0.041, respectively).
Acute Hormone Exposure
Thyroxine. Over the observation period of 6 hpa, the mean relative heart rate of
all treatment groups (0, 2.5, 5, 10 and 20 ng T 4/embryo) significantly increased in an
apparent linear fashion (F(3,
216)
= 87.479, p < 0.001; Tukey‟s, p‟s < 0.05; Fig. 2.4).
However, T4 administration of each doses (2.5, 5, 10 or 20 ng/embryo), did not
significantly alter this developmental trend among doses (p‟s > 0.05).
Triiodo-L-thyronine. There was a significant effect of T 3 dose (0, 2.5, 5 and 10
ng/embryo; F(3, 186) = 6.06, p < 0.001) and time (1, 2, 3 and 6 hpa; F (3, 186) = 73.708, p <
27
0.001) as well as an interaction of the two factors (F( 9, 186) = 3.065, p = 0.002) on mean
relative heart rate (Fig. 2.5).
Embryos exposed to T 3 doses of 0, 2.5, 5 and 10
ng/embryo, showed a significant time-dependent increase in mean relative heart rate
which, at 6 hpa, was significantly different between doses of 0 ng/embryo, and 5 and 10
ng/embryo (p‟s < 0.05). The mean relative heart rate of embryos administered 5 or 10
ng T3/embryo were approximately two-fold higher than that of embryos administered
vehicle alone.
Propyl-thiouracil. PTU had a significant dose-dependent effect (F(3, 203) = 3.96, p
= 0.009) on mean relative heart rate, but subsequent comparisons among doses (0, 15,
30 and 60 ug/embryo) revealed that PTU did not significantly alter mean relative heart
rate from vehicle-treated controls over the course of 6 hpa (Fig. 2.6). However, at 6
hpa, PTU treatment of 30 ug/embryo induced a significantly smaller increase in mean
relative heart rate than treatment of 60 ug/embryo (Tukey‟s, q = 4.06, p = 0.021). Mean
relative heart rate significantly increased over time among all dose groups (p‟s < 0.05)
except for the 30 ug/embryo dose group.
Testosterone. Over the 6 h period after TE administration, mean relative heart
rate significantly increased in a linear fashion for all dose groups (p‟s < 0.05), except for
the dose of 20 ng/embryo (Fig. 2.7). Administration of TE (2.5, 5, 10 and 20 ng/embryo)
at 70 h of age did not alter the magnitude of increase in mean relative heart rate from
the vehicle-treated group over the course of 6 h after administration.
Development Rate
The pharmacological agents used in this study did not significantly alter mean
HH stage from that of vehicle-treated controls for either administration ages (Figs. 2.8,
28
2.9, 2.10 and 2.11).Separate two-way ANOVAs for each pharmacological agent, with
respect to administration age (40 and 70 h) and dose revealed that the two
administration ages remained significantly different from one another during the course
of the experiment (T4: F(1, 204) = 197.08, p < 0.001; T3: F(1, 131) = 58.11, p < 0.001; PTU:
F(1, 85) = 244.86, p < 0.001; T: F(1, 103) = 219.48, p < 0.001).
Discussion
Heart Rate
Baseline Heart Rates
Baseline heart rate values were 13 to 28% higher than those embryos at similar
developmental stages in previous studies (Romanoff 1960; Clark and Hu, 1982;
Burggren et al., 2004). Baseline heart rate of chicken embryos is modestly increased
by development under the conditions of this experiment, perhaps because of altered
oxygen tension and humidity due to windowing.
Chronic Hormone Exposure
Administration of T3 to 40 or 70 h chicken embryos induced significant
tachycardia at 24 h post administration (Fig. 2.2).
This chronotropic response of
embryos was dose-dependent, and followed a sigmoidal dose-response relationship
that is common to physiological responses to toxicological and pharmacological agents
(Holford, 2005).
Accordingly, T 3 doses within the physiological range (2.5 and 5
ng/embryo) induced a gradual increase in heart rate as concentration increased; effect
of doses that extended beyond the physiological range (10 ng/embryo) was
characterized by a plateau (“ceiling effect”) of heart rate. This ceiling effect on heart
29
rate may reflect shock induced by toxicity, or the overshoot and/or undershoot of
homeostatic controls (Holford, 2005).
The cardiac response of early embryos to T3 detected was similar to that
observed in late stage embryos, juveniles and mature animals.
In adult and fetal
animals, hypothyroidism and the associated low levels of circulating thyroid hormones
induced via thyroid gland ablation resulted in smaller hearts and slower heart rates,
while hyperthyroidism had the opposite chronotropic effects (Balkman et al., 1992;
Lorijn et al., 1980). Although the cardiac response to perturbations in T 3 levels may be
similar in adult and developing animals (and in this case early developing animals), the
mechanisms that underlie the adult and embryonic responses are likely to be different.
For example, exposure to increased levels of T 3 throughout perinatal and postnatal
development increased cardiac alpha-adrenergic receptor density while T 3 exposure
during adulthood caused down-regulation of these receptors (Metz et al., 1996). Other
mechanisms that may explain the T 3-mediated tachycardia in developing animals
includes early maturation of cardiac myocytes (Chattergoon et al., 2007), and/or rapidly
changing pattern of thyroid hormone receptor expression throughout early development
(Sjöberg et al., 1992; Forrest et al., 1990). Unfortunately, these mechanisms are largely
unexplored in tissues of young embryos, and thus remain only speculations about T3mediated chronotropic effects observed in early developing chicken embryos of this
study.
Chronotropic response to T 4 detected in 64 h embryos were similar to the
responses to T4 reported for late stage embryos, juveniles and adults (Balkman et al.,
1992; Lorijn et al., 1980). Mechanisms that underlie these chronotropic responses are
30
likely to be similar to those suggested for T 3, since T4 activity in tissue is largely
characterized by its rapid conversion to T3 (Kuiper, 2005). Interestingly, heart rate of 94
h embryos treated at 70 h was relatively insensitive to doses of T 4 ranging from 2.5 – 20
ng/embryo, with T4 treated embryos showing a slight, but non-signficant decrease in
heart rate over vehicle-treated controls (Fig. 2.1). Perhaps thyroid hormone receptor
(TR) localization and density was rapidly changing during this early period of
development (40 – 94 h old), and causing heart rate to change in response to T 4
accordingly. Splicing of two genes, c-erbAα and c-erbAβ, in chickens, results in the
expression of two categories of TRs, TRα and TRβ, which are differentially expressed in
tissues during early development (Forrest et al., 1990; Sjöberg et al., 1992).
Additionally, the two types of TRs are functionally different; TRαs are directly involved in
heart contractibility, while TRβs are mainly involved in kidney and brain functions
(Chatterjee and Tata, 1992; Brent, 2000). At the onset of heart beat in chicken embryos
(~24 h), only c-erbAα transcripts are present in the embryo (Flamant and Samarut,
1998), suggesting that early development of the cardiovascular system may be
dependent upon TRα expression, while later in development (day 16), TRβs may also
be involved (Forrest et al., 1990; Mai et al., 2004). This differential expression of TRs
during development may be an explanation for the different actions of T4 at the two
developmental ages. Further investigations at the molecular and biochemical level are
necessary to assess whether receptor density and localization was responsible for the
age-dependent effects of T4. An alternative explanation is that the dose response curve
of T4 in 40 h embryos was significantly left-shifted in comparison to the dose response
curve of T4 at 70 h old, thus the complete dose-response curve of 70 h embryos was
31
not fully captured in this study. However this is unlikely given that T 3 increased heart
rate at this age at a dose of 5 ng/embryo, which is comparable in physiological potency
to T4 dose of 10 ng/embryo (Porterfield, 2001).
Although
age-dependent
physiological
and
morphological
responses
to
environmental and hormonal cues in developing animals is not a new concept (Metz et
al., 1996; Fritsch, 1997; Altimiras and Phu, 2000; Brent, 2000; Chan and Burggren,
2005), it is very interesting that an age difference of 30 h between the two early
developmental age groups was enough to capture a “critical window” in which two very
distinct chronotropic responses to T 4 occurred.
Equally interesting is that this
differential, age-dependent response to T4 was not detected in embryos exposed to
exogenous T3 (Fig. 2.2). T4 has traditionally been characterized as a less potent form of
T3, with physiological and morphological effects indistinctive from T 3 (Chopra et al.,
1978). Results reported here suggest that T4 and T3, although known collectively as
thyroid hormones, may act differently in early developing systems.
The rapid conversion of T 4 to the more potent form, T 3, is accomplished by the
cleaving of an iodine from T 4 via the action of iodothyronine-5‟-deiodinases type I and
type II (5‟DI and 5‟DII; Darras, 2006). These iodothryonine-5‟-deiodinases are active in
chicken embryos as early as 5 days old, and in particular, 5‟DI are detectable in the
hearts of chicken embryos during the last week of incubation (Van der Geyten et al.,
1997; Van der Geyten et al., 2002).
The present study showed that PTU, a
noncompetitive inhibitor of 5‟DI, had minimal effect on heart rates of early-stage
embryos, whether the administration occurred at 40 h or 70 h of age (Fig 2.3). PTU did
not have the effect that one would expect based on literature about juvenile and adult
32
animals. In mature animals, increase in T 3 and T4 plasma levels induces sustained
tachycardia, whereas PTU causes the opposite/antagonistic effect (Fazio, 2004;
Koenig, 2005; Kisso et al., 2008).
PTU has traditionally been employed to induce
hypothyroidism in animals with fully functionally thyroids, and have been used to
antagonize effects of thyroid hormones in in vitro experiments (Ma et al., 2003; Kisso et
al., 2008). In the current in vivo experiments, embryos lacked fully functional thyroid
glands, and were dependent on the maternally-derived T4 and T3 stores in the yolk. The
current observations suggest that, due to the large yolk T3 supplies available to the
embryo, 5‟DI activity is not a significant factor in modulating heart rate during early
development.
Testosterone is associated with increased risk of cardiovascular disease in adult
humans, partially due to its role in modulating contractility of cardiaomyocytes by
increasing calcium density in these cells (Golden, 2005; Er et al., 2007).
TE also
modulates early cardiac-specific, developmental events such as promoting the
differentiation of embryonic stem cell into cardiomyocytes (Goldman-Johnson et al.,
2008). However, the chronotropic effects of TE in developing animals remain poorly
understood.
In the current study, TE at doses of 2.5, 5, 10 and 20 did not affect
embryonic heart rates, when applied at either 40 h or 70 h of age (Fig. 2.3). However,
the non-significant increase in heart rate of embryos dosed at 70 h, and the slight, nonsignificant decrease in heart rate of embryos dosed at 40 h in concert caused the two
age groups to have significantly different heart rates at TE doses of 2.5 and 5
ng/embryo.
Since TE has been implicated in inhibiting the production of vascular
endothelial growth factor production from adult stem cells, and increasing contractility of
33
adult cardiomyocytes in vitro (Golden et al., 2005; Ray et al., 2008), perhaps the
observed embryonic chronotropic effects in this study result from these same
cardioregulatory mechanisms. However, there is no known evidence that TE induces
the same cardiac effects in embryonic cardiomyocytes, in vivo, as it does in vitro in adult
cardiomyocytes. Further understanding of the biochemical and cellular mechanisms of
TE‟s effects in embryonic cardiomyocytes at various points in development is necessary
to understand ultimately how TE induces an age-dependent chronotropic effect in early
development.
Acute hormone exposure
Another dimension of this study involved acute (1 to 6 h) effects of T3, T4, PTU
and TE on heart rates of early developing chicken embryos.
Chronic effects of
hormones such as the thyroid complex often involve receptor activation which leads to
changes in gene expression. Acute effects, however, are likely due to nongenomic
activity of the hormones, which includes activation of ion pumps and channels, and/or
interaction with signal transduction pathways to alter the intracellular ionic milieu
(D‟Arezzo et al., 2004; Davis, 2008).
Results of this study show that both T 3 and PTU (Figs. 2.5 and 2.6) had relatively
rapid chronotropic effects in 40 h embryos (6 hpa), while T 4 and TE did not (Figs. 2.4
and 2.7). T3 administration increased embryonic heart rate over the course of 6 hpa,
beyond the increase seen in vehicle-treated embryos.
This effect was observed
throughout the 6 h window, but reached statistical significance only at doses of 5 and 10
ng/embryo during the 6th hour of observation. Schwartz and Gordon (1975) showed
that T3 had no acute effect on the contraction rate of cultured chicken embryo heart
34
cells, suggesting that the in ovo, acute chronotropic effects of T3 observed in the
present study are likely due to an interaction of physiological processes rather than a
direct action on cardiomyocytes. Interestingly, PTU had no chronic chronotropic effects
in early embryos, even though PTU dampened the increase in heart rate at 6 hpa. The
effect occurred at a dose of 30 ug/embryo, and was significantly different from heart
rates heart rates of embryos given 60 ug/embryo (Fig. 2.6). This finding supports the
contention that chronic effects of some hormones are often very distinct from their acute
effects (Er et al., 2007).
Although TE and T4 have significant acute effects on in vitro adult and fetal
cardiac cells (Huang et al., 1999; Er et al., 2007), no short-term chronotropic effects of
TE or T4 occurred during early development, in vivo. Overall, these data suggest that
embryonic cardiac cells respond differently from adult cardiac cells to acute
perturbations in T4 and TE levels, and that in vitro responses of cardiac cells to
hormonal perturbations are entirely different from in vivo cardioregulatory responses in
early development. Further biochemical and cellular investigations are necessary to
clarify the interpretation of these observations.
Development Rate
Development rates of 40 h and 70 h embryos did not differ significantly among
the treatment groups for each of the pharmacological agents (Figs. 2.8- 2.11). At each
dose of T3, T4, TE, and PTU, the frequency of embryos at each developmental stage did
not vary from vehicle or the other doses. Heart rate was significantly altered 24 h after
administration by T3 applied at 40 h and 72 h, and by T 4 applied at 40 h, yet
development rate (based on gross morphology) in these embryos remained unaltered
35
from vehicle-treated embryos.
Collectively, these data show that during early
development, development rate was not significantly affected by perturbations in yolk
hormone levels. Therefore, chronotropic effects of these hormones (T3, T4 and TE)
were not a result of acceleration of development rate. Cardiac activity in early chicken
embryos (3-4 days incubation) does not play a role in overall growth or oxygen
consumption; rather, it potentially modulates/regulates cardiac maturation and
angiogenesis via hemodynamic (blood flow and pressure dynamics) action (Burggren et
al., 2000; Burggren, 2004). Thus, it is reasonable that hormones that induced changes
in heart rate did not necessarily induce comparable changes in overall development
rate.
Summary
This study provides the first comprehensive study of the chronotropic effects of
acute and chronic exposure to physiologically-relevant hormones at two distinct periods
of early embryonic development of the chicken.
I have shown that modulation of
hormones that are known to exist in the egg yolk at physiologically significant
concentrations can elicit very different age-dependent cardiac responses in the early
developing chicken embryo. In light of evidence that these hormones exist under highly
variable levels in egg yolks within populations, within species and among species, it is
important to understand how these maternally-derived factors affect early embryonic
development. The next two chapters assess the significance of inter-species and intraspecies variation in naturally-occurring yolk hormones (T3 and TE) in the modulation of
the early embryonic phenotypes of avian embryos.
36
15
(28)
10
fH Change (%)
*
*
5
(25)
(25)
(29)
(81)
0
(26)
(10)
-5
(10)
(11)
(11)
-10
0
5
10
15
20
T4 Concentration (ng/embryo)
FIGURE 2.1 Mean relative heart rate (recorded 24 hpa) of chicken embryos exposed
to T4 (0, 2.5, 5, 10 or 20 ng/embryo) at 40 h (unfilled triangles), or at 70 h of age (filled
triangles). A line (dashed = 70 h; solid = 40 h) indicates no difference among included
dose groups within a given administration age. An asterisk indicates a significant
difference between the two administration ages at the given dose. N values for each
mean value are indicated in parentheses. Data are presented as mean + S.E.M.
37
15
(11)
fH Change (%)
10
(7)
(6)
5
(15)
(25)
(7)
(12)
0
(13)
-5
-10
0.0
2.5
5.0
7.5
10.0
T3 Concentration (ng/embryo)
FIGURE 2.2 Mean relative heart rate (recorded 24 hpa) of chicken embryos exposed to
T3 (0, 2.5, 5 and 10 ng/embryo) at 40 h (unfilled squares), or at 70 h of age (filled
squares). A line (dashed = 70 h, solid = 40 h) indicates no difference among included
dose groups within a given administration age. N values for each mean value are
indicated in parentheses. Data are presented as mean + S.E.M.
38
15
10
*
fH Change (%)
(10)
5
*
(11)
(10)
(10)
(14)
0
(15)
-5
(8)
(7)
(8)
(6)
-10
0
5
10
15
20
TE Concentration (ng/embryo)
FIGURE 2.3 Mean relative heart rate (recorded 24 hpa) of chicken embryos exposed to
TE (0, 2.5, 5, 10 or 20 ng/embryo) at 40 h (unfilled diamonds), or at 70 h of age (filled
diamonds). A line (dashed = 70 h old, solid = 40 h old) indicates no difference among
included dose groups within a given administration age. An asterisk indicates a
significant difference between the two administration ages at the given dose. N values
for each mean value are indicated in parentheses. Data are presented as mean +
S.E.M.
39
fH Change (%)
30
0 ng / embryo (n>9)
2.5 ng / embryo (n>8)
5 ng / embryo (n>9)
10 ng / embry (n>8)
20 ng / embryo (n>8)
20
10
0
0
1
2
3
4
5
6
7
Time after administration (hours)
FIGURE 2.4 Mean percent change in heart rate over time of chicken embryos exposed
to T4 (0, 2.5, 5, 10 or 20 ng/embryo) at 40 h of age. A line indicates no difference
among included dose groups within a given time after administration. N values for each
mean value are indicated in parentheses. Data are presented as mean + S.E.M.
40
fH Change (%)
30
0 ng / embryo (n>13)
2.5 ng / embryo (n>7)
5 ng / embryo (n> 6)
10 ng / embryo (n>6)
20
10
0
0
1
2
3
4
5
6
7
Time after administration (hours)
FIGURE 2.5 Mean percent change in heart rate over time of chicken embryos exposed
to T3 (0, 2.5, 5 or 10 ng/embryo) at 40 h of age. A line indicates no difference among
included dose groups within a given time after administration. N values for each mean
value are indicated in parentheses. Data are presented as mean + S.E.M.
41
fH Change (%)
30
0 ug / embryo (n>7)
15 ug / embryo (n>10)
30 ug / embryo (n>10)
60 ug / embryo (n>10)
20
10
0
0
1
2
3
4
5
6
7
Time after administration (hours)
FIGURE 2.6 Mean percent change in heart rate over time of chicken embryos exposed
to PTU (0, 15, 30 or 60 ug/embryo) at 40 h of age. A line indicates no difference among
included dose groups within a given time after administration. N values for each mean
value are indicated in parentheses. Data are presented as mean + S.E.M.
42
fH Change ((%)
30
0 ng / embryo (n>9)
2.5 ng / embryo (n>9)
5 ng / embryo (n>9)
10 ng / embryo (n>9)
20 ng / embryo (n>9)
20
10
0
0
1
2
3
4
5
6
7
Time after administration (hours)
FIGURE 2.7 Mean percent change in heart rate over time of chicken embryos exposed
to TE (0, 2.5, 5, 10 and 20 ng/embryo) at 40 h of age. A line indicates no difference
among included dose groups within a given time after administration. N values for each
mean value are indicated in parentheses. Data are presented as mean + S.E.M.
43
Developmental Stage (HH)
22
21
20
(12)
(6)
(6)
(10)
(10)
19
18
17
16
*
*
(27)
*
*
(23)
(27)
5
10
*
(26)
(58)
15
14
0
0
15
20
T4 Concentration (ng/embryo)
FIGURE 2.8 Mean stage (recorded 24 hpa) of chicken embryos exposed to T 4 (0, 2.5,
5, 10 or 20 ng/embryo) at 40 h (unfilled triangles), or at 70 h of age (filled triangles). A
line (dashed = 70 h old, solid = 40 h) indicates no difference among included dose
groups within a given administration age. An asterisk indicates a significant difference
between the two administration ages at the given dose. N values for each mean value
are indicated in parentheses. Data are presented as mean + S.E.M.
44
Developmental Stage (HH)
22
21
20
19
(13)
(7)
(6)
(6)
18
17
16
*
*
(19)
(41)
*
*
(23)
(17)
15
14
0
0.0
2.5
5.0
7.5
10.0
T3 Concentration (ng/embryo)
FIGURE 2.9 Mean stage (recorded 24 hpa) of chicken embryos exposed to T 3 (0, 2.5,
5, 10 or 20 ng/embryo) at 40 h (unfilled squares), or at 70 h of age (filled squares). A
line (dashed = 70 h, solid = 40 h) indicates no difference among included dose groups
within a given administration age. An asterisk indicates a significant difference between
the two administration ages at the given dose. N values for each mean value are
indicated in parentheses. Data are presented as mean + S.E.M.
45
Developmental Stage (HH)
22
21
20
(9)
(11)
(15)
(8)
19
18
17
*
*
*
16
15
(17)
*
(10)
(8)
(8)
14
0
0
10
20
30
40
50
60
PTU Concentration (ug/embryo)
FIGURE 2.10 Mean stage (recorded 24 hpa) of chicken embryos exposed to PTU (0,
15, 30 or 60 µg / embryo) at 40 h (unfilled circles), or at 70 h of age (filled circles). A
line (dashed = 70 h, solid = 40 h) indicates no difference among included dose groups
within a given administration age. An asterisk indicates a significant difference between
the two administration ages at the given dose. N values for each mean value are
indicated in parentheses. Data are presented as mean + S.E.M.
46
Developmental Stage (HH)
22
21
20
19
(17)
(14)
*
*
(11)
(10)
(11)
18
17
16
15
(13)
*
*
*
(8)
(7)
(6)
5
10
(7)
14
0
0
15
20
TE Concentration (ng/embryo)
FIGURE 2.11 Mean stage (recorded 24 hpa) of chicken embryos exposed to TE (0,
2.5, 5, 10 or 20 ng/embryo) at 40 h (unfilled diamonds), or at 70 h of age (filled
diamonds). A line (dashed = 70 h, solid = 40 h) indicates no difference among included
dose groups within a given administration age. An asterisk indicates a significant
difference between the two administration age at the given dose. N values for each
mean value are indicated in parentheses. Data are presented as mean + S.E.M.
47
CHAPTER 3
INFLUENCE OF INTER-SPECIES AVIAN YOLK FACTORS ON HEART RATE,
OXYGEN CONSUMPTION AND BODY MASS OF
EARLY-STAGE CHICKEN EMBRYOS
Introduction
Adult phenotypes among bird species show marked differences that can be
traced back to differences in developmental patterns during embryonic stages of life
(Anthony et al., 1991; Lilja et al., 2001; Blom and Lilja, 2005).
Interestingly, many
embryonic traits (i.e. length of prenatal development, body mass, metabolic rate and
heart rate) that significantly vary among bird species scale with the egg mass (Rahn et
al., 1974; Rahn and Ar, 1974; Tazawa et al., 1991; Deeming et al., 2006; Deeming and
Birchard, 2007).
For example, species-specific differences in length of prenatal
development among avian species are characterized by a wide range of lengths of
incubation periods (from 13 days in quail to 41 days in ostrich) which allometrically scale
with species-specific egg masses (from 11 g in quail to 1400 g in ostrich; Rahn and Ar,
1974). Also, during the last quarter of embryonic development, heart rates of nearpipped (late-stage) bird embryos inversely scale with inter-species egg mass (Tazawa
et al, 1991), while oxygen consumption of the egg is directly related to egg mass (Rahn
et al, 1974). Moreover, among bird species, initial egg mass strongly predicts hatchling
mass (Deeming and Birchard, 2007).
Egg shell thickness is allometrically related to egg mass, thus the conductance
(or diffusivity to oxygen) of the egg shell generally decreases as egg size increases (Ar
et al., 1979). This phenomenon has been suggested to be a major predictor of avian
48
physiology such as developmental rate, heart rate, oxygen consumption, and growth
rate late in embryonic development (Rahn et al, 1974; Rahn and Paganelli et al, 1982;
Ar and Rahn, 1985; Rahn et al, 1987; Meir and Ar, 1990). However, it is not clear if and
how early-stage embryonic physiological patterns scale with egg mass among bird
species. Romanoff (1960) suggests that heart rate of three bird species (quail, chicken
and duck) during the first quarter of incubation may be inversely correlated with egg
mass.
This heart rate – egg mass relationship is similar to that observed in late
incubation stages (Rahn et al, 1974; Ar and Tazawa, 1999; Tazawa et al., 2001). In
contrast, the relationship between egg mass and development rate of late-stage
embryos does not necessarily apply to the early stages of embryonic development
(Sellier et al., 2006). Thus it is of interest to investigate whether egg mass can predict
inter-species physiological patterns of early-stage embryos in the manner that it does
during the late stages of embryonic development.
Bird eggs possess a wide range of maternally-derived yolk factors that can
potentially alter early embryonic heart rate and general embryonic physiology, and
growth rate (Schwabl, 1993; Wilson and McNabb, 1997; Badzinski et al., 2002; Gorman
and Williams, 2005; see also results of Chapter 2). Thus it is possible that the heart
rate – egg mass relationship among late-stage embryos of bird species can be
explained by the effects of maternally-derived yolk factors that covary with egg mass
and/or length of incubation period early in embryonic development. Supporting this
conjecture is the strong inverse relationship between prenatal development length (as
indicated by length of incubation period) and yolk testosterone (TE) concentrations in
the egg yolk of passerine bird species (Gorman and Williams, 2005). As of yet, the
49
covariation of yolk hormones and incubation period or egg mass is poorly understood in
non-passerine, precocial species.
Moreover, the physiological significance of inter-
species variations in yolk hormone concentrations has not been experimentally
assessed in early developmental periods (i.e. the first few days of avian embryonic
development).
I hypothesized that egg mass and length of incubation period of non-passerine,
precocial bird species can predict the concentrations of triiodothryonine (T 3) and
testosterone (TE) in the egg yolk on an inter-species level. To test the hypothesis, T 3
and TE radioimmunoassay were carried out on the egg yolks of 7 species from 2
parvclasses (Ratitae which includes the order Struthioniformes; and Galloanserae which
includes the orders Galliformes and Anseriformes) of the subclass Neornithes.
Also, I hypothesized that exposing chicken embryos to the yolks of different bird
species causes significant and predictable changes in early embryonic heart rate, massspecific oxygen consumption and body mass, and that the phenotypic changes scale
with egg mass, length of incubation period and yolk hormone concentrations that is
representative of the bird species from which the yolk environment was derived.
A
technique by which to completely remove chicken embryos from their original yolk and
transfer them to the yolks of other bird species during the first few days of embryonic
development was developed to test this hypothesis.
Materials and Methods
Egg Storage and Incubation
White Leghorn chicken eggs for incubation were purchased from Texas A&M
University (College Station, TX), while eggs for yolk analyses and manipulation were
50
purchased from a variety of small, independent farms and businesses from across the
United States. Upon arrival, eggs were stored at room temperature (23-26°C) for a
maximum of 5 days until the start of incubation or yolk collection. Chicken eggs were
incubated in commercial incubators at 37°C and 58% humidity, and were automatically
turned every hour during the course of the incubation period. Humidity and temperature
were checked daily.
Experimental Design
This study consisted of two experiments. The first experiment was undertaken to
characterize the inter-species relationship between egg mass and yolk hormones,
incubation period and yolk hormones, as well as egg mass and incubation period. T 3
and TE radioimmunoassays were performed on the yolks of a variety of non-passerine,
precocial avian species (subclass: Neornithes). Species under investigation included
common quail (Coturnix coturnix), chicken (Gallus gallus, breeds: White Leghorn,
Cornish Rock, Buff Cochin, Buff Cochin Bantam, South Carolina White Leghorn
Bantam), duck (Anas platyrhynchos, breeds:
Mallard, Khaki Campbell, Blue Indian
Runner, and Buff Runner), Narragansett turkey (Meleagris gallopavo), Chinese goose
(Anser cygnoides), emu (Dromaius novaehollandiae), and ostrich (Struthio camelus).
For each species, except for ostrich and emu, eggs were obtained from at least two
different suppliers to reduce the potential confounding environmental factors such as
feed type and climate, and biological factors such as maternal condition, on egg and
offspring quality (Latour, 1996; Rozenboim et al, 2007). For regression analyses, yolk
hormone concentrations and egg masses were experimentally determined, while
51
incubation period was derived by obtaining the mean value of the shortest and longest
reported incubation period for each species.
The second part of this study involved growing chicken embryos on the yolk of
other species to assess how species-specific variation in yolk factors influences
embryonic physiology and morphology during early development. Chicken embryos
were removed from their original yolk after 40 hours (h) of in ovo incubation, and placed
on culture dishes filled with one of a variety of yolks derived from the eggs of the
majority of the avian species mentioned above. Chicken embryos developed for an
additional 36 h on the foreign yolks, after which, heart rate (f H), mass-specific oxygen
consumption (
),
and wet body mass (BM) were assessed.
At the time of
assessment, embryos were ~3 days old. Physiological and morphological data were
regressed with initial egg mass (IEM), incubation period (IP), and yolk T 3 and yolk TE
concentration of the egg from which yolks were derived to assess if species-specific egg
mass, length of prenatal development, and yolk environment can significantly predict
embryonic phenotypes in a species-specific manner.
It is unconventional to suggest that heart rate varies as a function of length of
prenatal development (incubation period).
However for the purpose of this study,
incubation period was viewed as a discrete characteristic of bird species (i.e. quail =
16.5 days, chicken = 21 days, etc.). Thus the regression of heart rate and incubation
period was performed to describe the possible link between yolk factors that covary with
incubation period, and their effects on heart rate.
52
Triiodo-L-thyronine Radioimmunoassay
Fresh,
unincubated
eggs were
dissected,
homogenized and stored at -20°C for assays.
and
yolks were
separated,
The methanol/ chloroform extraction
procedure of Wilson and McNabb (1997) was slightly modified to increase T 3 extraction
efficiency. 0.5 g of previously homogenized yolk was placed in 13ml glass conical tube
and 2ml of methanol (supplemented with 1mM propylthiouracil (PTU)) was added. The
components were mixed (via vortex mixer) for approximately 5 minutes to ensure a
homogenous suspension. Labeled thyroid hormone was added to each sample (~2000
cpm of
125
I-T4 or
125
I-T3; MP Biomedicals, New York, NY), mixed and counted using a
gamma counter. Subsequently samples were shaken (150 oscillations/min on a shaker)
for 10 min., and centrifuged for 10 minutes at 2700 rpm. The supernatant was decanted
into a 13ml glass conical tube and the precipitate was resuspended in 1ml methanol
(1mM PTU) and shaken, spun and decanted (as above) in another 13ml glass conical
tube. Five ml of chloroform and 0.5 ml of 2N ammonium hydroxide was added to each
tube, and the tubes were shaken and centrifuged. The upper phase of corresponding
tubes were removed and combined in a 13ml glass tube. The suspension was allowed
to dry overnight under a filtered air stream. The dried samples were resuspended in 1
ml 2N ammonium hydroxide, shaken, centrifuged and decanted into a 13ml tube. One
ml of chloroform was added to each tube, and the tubes were shaken, centrifuged, and
the upper phase was removed and allowed to dry under a filtered air stream. The
samples were resuspended in 150 µl phosphate buffered saline (supplemented with
alizarin) and counted (1 min). The extraction efficiency was calculated for each sample
53
by calculating the percentage of radioactivity that remained after the extraction
procedure.
T3 concentrations of the resuspended sample (from the last step of extraction
above) were analyzed using competitive binding radioimmunoassay kits (Free T 3
Antibody Coated Tube-125I RIA Kit, MP Biomedicals, New York, NY). Procedures of
the hormone kit were followed exactly.
Testosterone Radioimmunoassay
Testosterone concentrations in the previously collected yolk samples were
determined via radioimmunoassay following the protocol for steroid hormones in egg
yolks established by Schwabl (1993) and modified for use with quail, chicken, duck,
goose, turkey, emu and ostrich yolks. The protocol was modified by using a phosphate
buffered saline containing yolk stripped of hormones as the background buffer for the
standard curves. Using yolk pooled from eggs collected from all species controlled for
extra yolk material present in samples via direct assays. This negated the need for the
use of column chromatography to clean yolk samples. The protocol for stripping
hormones from yolk samples was modified from that of Wingfield et al. (1984) for
stripping plasma of steroids.
Egg Mass
Egg mass expressed in g was obtained from intact eggs prior to the use in
embryo culture experiments.
Embryo Cultures
Chicken embryos (~40 hours of incubation; Hamburger and Hamilton (1951)
stages 9-14) were removed from their original yolks and transferred to parafilm culture
54
dishes filled with the yolk of chicken, quail, duck, goose, turkey, emu or ostrich eggs by
following the methods previously outlined by Chapman et al (2001). Parafilm culture
dishes were prepared as previously described by Chapman et al. (2001), with the
exception of the culture medium used.
Parafilm culture dishes were prepared by
covering the lids of 35mm petri-dishes with parafilm, and completely filling them with
Yolk-Tyrode (YT) medium through a square opening (2 cm x 2cm) cut out of the
parafilm (Fig. 3.1). YT medium consisted of one part fresh, homogenized yolk from
layer or broiler eggs, and one part 1X Tyrode‟s Solution supplemented with 5 units
penicillin-streptomycin/ml (v/v; Sigma T1788, St. Louis, MO).
Embryos (HH 9-14) and associated vitelline membranes were carefully excised
from their native yolk using Whatman filter paper rings as the carrier. Embryo removal
from the yolk sac consisted of first removing albumen from the area surrounding the
embryo. Then a filter paper ring (outer diameter = 38 mm; inner diameter = 15 mm)
was positioned on the vitelline membranes so that the embryo was centrally framed by
the ring, and the surrounding membranes tightly adhered to the filter paper ring. The
membrane along the outer edge of the filter paper ring was cut and the paper ring
carrying the embryo was lifted from the yolk using fine-tipped forceps. Embryos were
washed free of residual yolk and albumen by gently swirling in three washes of warm 1X
Tyrode‟s, penicillin/streptomycin. The edge of the filter paper ring carrying the embryo
was blotted on a Kimwipe for a few seconds to remove excess liquid from the embryo,
and a second filter paper ring (outer diameter = 38 mm, inner diameter = 30 mm) was
placed on top of the embryo/filter paper as to sandwich the embryo and associated
vitelline membranes between the two filter paper rings (Fig. 3.1).
55
This entire
embryo/filter paper complex was then placed onto prepared parafilm culture dishes, with
the embryo dorsal side down, and in direct contact with the YT medium at the opening
in the parafilm dish.
Seven parafilm embryo cultures were placed into a large petri dish (135 mm X
10mm), which then was placed into a seal-top plastic chamber (170 mm diameter, 80
mm height) outfitted with inflow and outflow ports on the lid. Two Kimwipes, wetted with
10 ml of distilled water, were placed at the bottom of the plastic chamber to maintain a
high level of humidity. After the lids were placed on the containers, the containers were
flushed via the ports for 20 minutes with a gas mixture of 95% oxygen (O 2) and 5%
carbon dioxide (CO2; Wösthoff Gas Mixer, Calibrated Instruments, Inc., NY).
After
flushing of chambers, ports were sealed, and the whole container was maintained in a
37.5 + 0.2°C, clear-top Styrofoam incubator (Circulated Air Hova-Bator Incubator, Model
No. 1590, GA). The start of experiments was considered to be 1.5 hours after the
chambers have been sealed and placed in incubators.
The high oxygen atmosphere (95% O 2, 5% CO2) provided for the cultures in this
study was to increase survivability and proper development of cultured embryos.
In
vitro cell cultures are commonly maintained under this artificial environment to prevent
acidification of culture medium (personal communication with Susan Chapman). Heart
rate of culture embryos incubated under this atmosphere was similar to that of
windowed embryos (Chapter 2), and embryonic development progressed normally
according to Hamburger and Hamilton (1950).
The actual percent oxygen in the
chambers declined over the period of parafilm embryo culture incubation. Initial reading
was 90+5% O2, and decreased to 47+8% O2 after 24 h of incubation in a cohort of 5
56
chambers tested. For each trial, variability in the rate of PO 2 decline among chambers
was accounted for by evenly distributing experimental groups among all chambers.
Wet Body Mass
Live embryos were anesthetized via exposure to -20°C for 4 min. Fresh embryo
mass was determined by completely excising the embryo from the surrounding vitelline
membranes in 1X Tyrode‟s solution.
Embryos were then carefully blotted with
Whatmann filter paper for 20 seconds to remove all visible water surrounding the
embryo, and weighed using a microbalance.
Heart Rate
The heart beat of embryos maintained in clear-top incubator (Circulated Air
Hova-Bator Incubator, Model No. 1590, GA) set at 37.5+°C was visually counted over
the course of one min. Heart rates (fH) were reported as beats per minute (bpm).
Mass-Specific Metabolic Rate
Oxygen consumption of embryos was assessed by closed-system respirometry.
Individual metabolic respirometers were constructed from 118 ml mason jars with metal
screw-top lids. The bottom of the respirometer was filled with polyester casting resin to
reduce the respirometer air volume to 27 ml. The lid was outfitted with a syringe port
constructed from the hub of a 25 gauge needle to allow for the attachment of a metal
stopcock and a 5ml glass syringe (Proper MFG Co., Inc.) for air sampling. Individual
embryo cultures were placed in the center of each respirometer, and the lids were
tightly sealed to the mouth of the respirometer with the aid of petroleum jelly. The
respirometers were then placed into an acrylic reptile incubator set at at 37.5°C. Each
57
embryo in its respirometer was allowed to equilibrate for 30 min with the syringe port
open for free exchange of air between the respirometer and the ambient air.
Thirty min after placement of respirometers into the reptile incubator, a small 3way metal stopcock attached to a 5ml-glass syringe was inserted into the syringe port of
the lid, and a 1.5 ml sample of air from the respirometer was drawn into the syringe and
injected into a flow-through oxygen electrode (Model 16-730 Microelectrodes, Inc., NH)
to obtain the initial PO2 of gas in the respirometer. The voltage output of the electrode
was converted into partial pressure by Chart software (Power Lab:
Chart 5,
ADInstruments). The stopcock-syringe was returned to the port of the respirometers,
and the respirometers were sealed by setting the stopcock to allow passage of air
between the respirometer and the syringe only. After 60 min, 1.5 ml of respirometer gas
was drawn into the syringe, and the gas in the syringe was injected into the flow-through
oxygen electrode to obtain the final PO2 value (as described above). The mass-specific
oxygen consumption rate (
in mlO2.g.h-1) of transfer embryos was calculated using
the following equation:
Where ∆PO2 is the change in oxygen pressure of the gas over time interval (mmHg), v is
total volume of air in the respirometer (ml), Δt is the time elapsed between PO2 readings
(h), and m is the wet mass (g) of the embryo. Total gas volume was determined for
each metabolic respirometer by subtracting the weight of the empty respirometer from
the weight of the chamber filled with distilled water. An additional 5 g was subtracted
from the final weight to account for the volume taken up by the parafilm culture dish,
assuming that 1 g of distilled water occupied a volume of 1 ml.
58
Respirometers
containing yolk culture dishes without embryos were sampled during each trial run to
account for non-biological fluctuations in oxygen concentration of respirometer gas.
Mass-specific
of embryos was adjusted accordingly.
Statistical Analyses
In the first experiment, least-square regression analyses on log10-tranformed data
were performed to assess the relationship between factors of initial egg mass (IEM),
incubation period (IP), yolk T 3 concentration, and yolk TE concentration. Species of the
parvclass Ratitae investigated in this study (emu and ostrich) are highly distinct from
species of the parvclass Galloanserae in terms of thyroid hormone status and control
(Dawson, 1996). It was based on this that regression analyses were performed on
complete data sets (Neornithes), and data subsets of Galloanserae and Ratitae. This
allowed for the assessment of taxon-specific differences among Neornithes. Due to the
small number of Ratitae species under investigation, regression analyses were
interpreted conservatively for this taxon.
In the second experiment, regression analyses were performed on a combination
of factors (independent factors: IEM, IP, yolk T3, and yolk TE; dependent factors: wet
body mass (BM), mass-specific oxygen consumption (
), heart rate (fH), and stage
(HH) to determine how much of the variation in physiological and morphological traits of
chicken embryos cultured on the different avian yolks can be accounted for by the
variation in the species-specific egg components and IP.
As discussed above,
regression analyses were performed on the complete data set, as well as a subset of
data which excluded the Ratitae. In some cases factors did not lend themselves to a
regression of raw data due to the design of the experiment (i.e. yolks were combined to
59
make culture medium; and yolk T 3 and yolk TE were assessed in eggs different from
those used for Embryo Culture experiments).
In these instances, the independent
factor data was first averaged within each species (i.e. mean yolk T3 concentration for
each species), and then the average values were regressed with the dependent factor.
For all statistical analyses, statistical significance was determined by P < 0.05.
Regression models are presented as y = axb(SEM). Values were presented as mean +
SEM.
Here, I used a species-correlation analyses approach, thus assuming that the
inter-specific data are phylogenetically independent. Since my goal was to investigate
the variability among inter-species traits in the current environment, and how this
current-day variability in traits among avian species impacts chicken embryo physiology
(Price 1997), I chose not to control for phylogenetic relatedness (i.e. independent
contrast analyses) in the statistical analyses.
Results
Yolk hormones, egg mass and incubation period in Neornithes
Regression of incubation period and initial egg mass
Table 3.1 shows the descriptive statistics of the factors on which least square
regression analyses were performed. Initial egg mass for 7 species from 3 orders of
birds ranged from an average of 1660.5 g in ostrich to merely 11.9 g in common quail,
while incubation period ranging from 56 days in emus down to 16.5 days in common
quail (Table 3.1).
To characterize the nature, strength and significance of the
relationship between IP and IEM within subclass (Neornithes), within parvclass
(Galloanserae) and within order (Galliforme), IP was regressed on IEM based on all
60
species data (Neornithes), and subsequently on only Galloanserae and Galliformes
(Fig. 3.1). Least-square regression analyses showed a strong and highly significant
relationship between IP and IEM at all three taxanomic levels (Fig.3.2).
Statistical
comparison of the three regressions indicated that the slopes (b) and intercepts (a) of
the regression models of subclass and parvclass were not significantly different from
each other.
However, the slope of the regression of order was highly significantly
different from both that of the regression of subclass (F (1, 206) = 23.51, p < 0.0001) and
that of the regression of parvclass (F (1, 90) = 13.34, p = 0.0003). This strongly indicated
that as initial egg mass among bird species increased, incubation period followed suite,
but that the nature of the relationship (defined by the slope of the regression) was
taxon-specific.
Regression of yolk hormones and initial egg mass
Yolk T3 concentrations among species ranged from ~0.46 to 3.16 pg mg -1, while
yolk TE ranged from 2.28 to 5.62 pg mg -1 (Table 3.1). Among Neornithes, yolk T3 or
yolk TE did not vary with IEM in a predictable manner (Table 3.2 and Fig. 3.3A and B).
However when Neornithes were parsed according to parvclass, IEM was significantly
related to yolk T3 and yolk TE in a parvclass-specific manner. In other words, with
respect to the eggs of Galloanserae (quail, chicken, turkey, duck and goose), yolk T 3
increased as IEM increased (yolk T3 = -0.597 IEM 0.409(0.155), r2 = 0.145, p = 0.012), while
for Ratitae eggs, yolk T3 and IEM were significantly inversely related (r2 = 0.816, p =
0.014). It must be noted, however, that the large egg mass large difference between
ostrich and emu species was likely to exaggerate the strength and direction of this
relationship.
61
Similar to yolk T3, yolk TE changed in direct relation to IEM among Galloanserae
(r2 = 0.066, p = 0.006); however no relationship was detected between the two factors
among the eggs of Ratitae (Table 3.2 and Figure 3.3B). Overall, IEM accounted for a
relatively small, but significant, percentage of the variation in yolk T 3 and yolk TE of
Galloanserae (r2 = 0.066, p = 0.006).
Regression of incubation period and yolk hormones
Yolk T3 concentration of the eggs of Neornithes was significantly related to
incubation period of the species to which the egg belongs (Table 3.3 and Fig. 3.4A).
The removal of Ratitae from regression analyses, did not alter the relationship between
yolk T3 and IP (p > 0.08). This suggests that the nature of the relationship between IP
and yolk T3 was not specific to taxonomic grouping. In contrast to yolk T3, it appeared
that yolk TE predicted IP in a taxon-specific manner. IP significantly increased as yolk
TE increased among Galloanserae only (Fig. 3.4B; IP = 1.29 yolk TE0.10(0.03), r2 = 0.07, p
= 0.005).
Moreover, within Galloanserae, yolk T 3 accounted for nearly 30% of the
measured variation in IP, while yolk TE only explained 6.7% of the variation in IP (Table
3.3). This difference indicated that yolk T 3 explained the variation in incubation period
among species better than yolk TE.
Modulation of Chicken Embryo Phenotype by Different Yolks
Factors in the prediction of body mass
Table 3.4 lists the means + S.E.M. of the morphological and physiological traits
of chicken embryos that were exposed to yolks belonging to a variety of precocial avian
species. The initial mass, yolk hormones and incubation period of these eggs were
previously assessed in the first experiment of the study. The relationship between BM
62
of chicken embryos and yolk T 3, log yolk TE, log IEM and log IP of the yolk on which the
chicken embryos were cultured are depicted in Figure 3.5A, B, C and D. Comparisons
of slopes and intercepts of least-square regressions of BM and corresponding factors,
based on the inclusion or exclusion of Ratitae, revealed that the exclusion of this
parvclass significantly modified the regression models with independent factors of IEM
(Fig. 3.5C), IP (Fig. 3.5D) and yolk TE (Fig. 3.5A) but not the model with independent
factor of yolk T3 (Fig. 3.5B).
When parvclass Ratitae was removed from the regression model of BM and IEM
(Fig. 3.4C), IEM became a significant predictor of chicken embryo mass (BM = 0.885
IEM0.10(0.04), r2 = 0.071, p = 0.015). Similarly, IP was significantly related to BM when
Ratitae were excluded from the model (BM = 0.387 IP 0.49(0.162), r2 = 0.102, p = 0.003).
As for the relationship between BM and yolk TE, the regression models including and
excluding Ratitae were both significiant (BM = 0.87 yolk TE 0.33(0.11), r2 = 0.07, p = 0.004,
and BM = 0.63 yolk TE0.70(0.18), r2 = 0.16, p <0.001, respectively), and the slopes of the
two models were significantly different from one another (F (1, 193) = 4.09, p = 0.04). In
contrast, the inclusion or exclusion of Ratitae did not significantly change the slope or
intercept of the significant allometric relationship between BM and yolk T 3 (slope
comparisons, p = 0.69). Based on these results, I concluded that the initial egg mass
and incubation period of species that are of the parvclass Ratitae do not predict BM as
strongly as do the initial egg mass and incubation period of species that are of the
parvclass Galloanserae. However, the body mass of chicken embryos subjected to
different avian yolks during development can be reliably predicted from the yolk T 3 and
63
yolk TE concentrations of the those yolks, regardless of the parvclass to which they
belong.
Factors in the prediction of heart rate
The variation in heart rate of chicken embryos cultured on the yolks of
Galloanserae eggs was significantly accounted for by the yolk T 3 concentration (Fig.
3.5A), yolk TE concentration (Fig. 3.6B), initial egg mass (Fig. 3.6C), and incubation
period (Fig. 3.6D) of those eggs. The inclusion of Ratitae in the regression models
changed both the predictability of the model (r2) as well as the nature of the relationship
between the independent factors and f H, except for the relationship between yolk T 3 and
fH. For the factor of yolk TE, the removal of Ratitae from the regression model caused
the factor to become a significant predictor of fH (fH = 2.17 yolk TE0.11(0.04), r2 = 0.019, p =
0.02); however the model only accounted for ~2% of the variation in f H. Moreover, the
slopes of the regressions of IEM and f H were significantly different based on the
inclusion or exclusion of Ratitae. This was also true for regressions generated for IP
and fH (Fig. 3.6D). The parameters of Ratitae yolks prove to be different from those of
Galloanserae yolks in their ability to predictably change the heart rate of chicken
embryos, with the exception of yolk T 3.
Factors in the prediction of mass-specific
.
Relative to body mass and heart rate, few factors were significant predictors of
mass-specific
of cultured chicken embryos (Fig 3.7A-D). Only yolk TE (Fig. 3.7B)
and IEM (Fig. 3.7C) were significantly related to mass-specific
of chicken embryos
cultured on the egg yolks of Galloanserae, with yolk TE also showing a significant
inverse relationship with mass-specific
of chickens cultured on the yolks of
64
Neornithes as a whole (VO2 = 0.76 yolk TE-0.36(0.15)). Yolk T3, which was a significant
predictor of BM and fH, failed to account for the any variation measured in mass-specific
(Fig.3.7A). Furthermore, the models generated for yolk TE and
explained less than 6% of the variation in mass-specific
, and IEM and
of cultured chicken
embryos.
Discussion
Relationship among yolk hormones, egg mass and incubation period of Neornithes
The robust relationship between IEM and IP among and within superorders and
orders of Neornithes species detected in the present study corroborates earlier studies
of Rahn and Ar (1974). The exponents (b) obtained for the regression models for
Neornithes, Galloanserae and Galliformes were in the range ~0.23 (Fig. 3.2), which
coincides with Rahn and Ar (1974)‟s exponent of 0.22, and Ar and Tazawa‟s (1999)
exponent of 0.21.
The exponents derived for previous studies were criticized for not
taking into consideration the interdependence of inter-species data (Deeming and
Birchard, 2006). When phylogenetic relatedness was accounted for, the models which
predict the relationship of IEM and IP among avian species did not adequately account
for inter-order differences. In other words, one universal regression model cannot be
applied to all bird orders to reliably predict IP from IEM, but rather, separate models
must be generated for each order (Deeming et al, 2006). In the current study, the
significant change in the slopes when models were parsed based on superorder and
order supports this concept of differential relationship of IP and IEM based on
phylogenetic taxa (Fig. 3.2).
65
An important and novel finding of this study is that length of incubation period can
be predicted from yolk T 3 and yolk TE in non-passerine, precocial birds.
These
observations fill a gap in the literature pertaining to the identification of physiological
factors (i.e. yolk hormones), other than initial egg mass, that can be used to predict
incubation period (and thus developmental rate and growth rate) in non-passerine
species (Deeming et al, 2006). Among passerine species (all of which are altricial),
length of incubation period and yolk TE are inversely related (Gorman and Williams,
2005), while the current study of non-passerine, precocial species of Galloanserae
revealed that IP and yolk TE was defined by an direct relationship (Fig.3.3B). Precocial
birds display well-developed features and are capable of locomotion upon hatch, while
altricial birds are relatively less mature and require extensive parental care upon hatch
(Nice, 1962). Also, precocial, non-passerine birds (such as the quail) show marked
differences from the altricial birds (such as the fieldfare and ring dove) in the rate of
embryonic organogenesis and ontogeny of hormonal function (McNabb 1988, Blom and
Lilja, 2005). The reversal of the IP – yolk TE relationship between the two bird groups
may reflect the role of maternal effects in the observed embryonic morphological and
physiological differences between altricial and precocial species. Moreover, this study
demonstrated that the relationship between IP and yolk hormones was also different
within precocial species.
The parvclass Galloanserae showed IP – yolk hormone
relationships that were distinct from, or not detected in Ratitae (Fig. 3.4A, B). This
distinction in yolk hormone concentrations among parvclasses suggests that yolk
hormones may underlie the early embryonic differences that exist between
Galloanserae and Ratitae.
For example, the nature of embryonic cardiovascular
66
maturation in Galloanserae species, such as the chicken, and Ratitae species, such as
the emu, are largely different (Crossley et al., 2003). It is possible that the difference in
IP - yolk hormone relationship between the parvclasses drive the difference in the
ontogeny of cardiovascular control between chicken and emu.
Collectively, the findings of this study strongly suggest that phylogeny plays a
critical role in the predictability of IP as a function of deposition of yolk hormones into
the egg, and yolk hormones significantly vary with egg mass, and that the alteration in
this relationship may reflect the early embryonic differences observed among bird taxa
and species.
However, biologically-relevant factors other than egg mass and yolk
hormones are likely to be involved in explaining the variation in yolk hormone
concentrations and incubation period length in precocial eggs since r2 values were
relatively small.
Modulation of Chicken Embryo Phenotype by Different Yolks
The physiological significance of the relationship between yolk hormones and
egg mass/incubation period among bird species was tested in the second part of this
study.
Heart rate, mass-specific
and body mass of chicken embryos were
significantly altered when they were exposed to the yolks of different bird species during
the first few days of development (Table 3.4). There was a significant direct relationship
between the yolk-induced physiological changes in the chicken embryos and the
hormone concentrations of the yolk on which the embryos were cultured (Figs.3.5, 3.6
and 3.7) These findings strongly suggested that the relationship between yolk hormone
and egg mass, and yolk hormones and incubation period (demonstrated in first part of
67
this study) among precocial bird species are of physiological and morphological
significance during early periods of bird embryo development.
In Chapter 2, T3 dose-dependently increased the heart rate of 3-4 day old
chicken embryos, while TE had no chronotropic effects. In this study, increase in both
yolk T3 and yolk TE correlated with an increase in the heart rates of cultured chicken
embryos. The differential effects of yolk TE on embryonic heart rate between this study
and the study of Chapter 2 is likely due to the variation of other yolk factors among bird
species. Bird egg yolk contains a wide variety of physiologically-relevant constituents,
and these constituents vary dramatically among species (Groothuis et al., 2005). It may
be that the correlation of heart rate and yolk TE is due to the covariation of TE with
another yolk factor that affects heart rate. A more complete characterization of the
hormone profile of the yolks of these bird species would be helpful in the greater
understanding of the relationship between inter-species variation in yolk factors and the
physiological patterns induced by modifying yolk environment.
In comparison to previous studies that have investigated the inter-species
relationships between egg mass and physiological attributes of the embryo (i.e. heart
rate or metabolic rate) (Rahn et al., 1974; Sotherland and Rahn, 1987; Tazawa et al,
1991; Ar and Tazawa, 1999; Deeming and Birchard, 2006), the present study is unique
in two major respects: 1) developmental age at time of assessment and 2) experimental
approach. Previous studies demonstrating that egg mass significantly predicts heart
rate and metabolic rate within and among bird species have done so in late-stage
embryos (60-95% incubation; Rahn et al., 1974; Tazawa et al., 1991). The current
study is the first to characterize these relationships in early-stage embryos (~14%
68
incubation), when physiology is quite unique in comparison to late-stage physiology
(Burggren et al., 2000; Burggren et al., 2004; Romanoff 1960). I have demonstrated
that early in development, chicken embryos are significantly responsive to speciesspecific shifts in yolk environment, and that these yolk-induced responses are
accounted for by the egg mass and length of incubation period that is characteristic of
the bird species to which the yolk environment belongs. This strongly suggested that at
a time in embryonic development when heart rate, metabolic rate, and body mass are
largely independent of the direct consequences of egg mass, egg mass and incubation
period of a species covary with yolk factors that significantly influence embryonic heart
rate, body mass and metabolic rate. Possibly, maternally-derived yolk hormones that
play a role in regulating the length of prenatal development altered the physiology of the
chicken embryo as early as the first few days of embryonic development. The detection
of these relationships early in development is intriguing and warrants further
investigation.
In terms of experimental approach, previous studies characterized the
relationship between egg constituents and the phenotype of bird embryos in their
natural yolk environments (i.e. in ovo) (McNabb, 1988; Schwabl, 1993; Gorman and
Williams, 2005).
The current study is the first to assess the physiological and
morphological traits of embryos of a single species (the chicken), in the context of the
yolk environment of a wide range of species. By isolating the epigenetic effects of yolk
environment from the direct genetic effects of phylogeny, it was possible to assess the
influence of egg yolk, per se, on the early embryonic phenotypes observed among avian
species. An interesting finding of this study is that 16.2% of the variation in body mass
69
of early chicken embryos was accounted for by yolk TE concentration (and egg mass)
of the yolk environment (Fig. 3.5B).
This finding extends previous reports of the
influence of yolk TE on the growth of bird hatchlings to early stages of embryonic
development. In late-stage embryos, yolk TE affects body mass negatively, while an
increase in yolk TE during embryonic stages causes an increase in growth rate
postnatally (Rubolini et al., 2005; Schwabl, 1996). Collectively, these data suggests
that inter-specific differences in yolk TE, per se, affect growth of birds in an agedependent manner.
Unfortunately, not much is known of the early-stage embryonic heart rate and
metabolic rate of the bird species used in the study. To date only a handful of studies
have assessed heart rates and metabolic rates of 3-4 day old chicken embryos
(Romanoff, 1960; Burggren et al, 2000; Burggren et al, 2004). Even less studied is the
impact of yolk hormones on these early embryonic traits and how these particular
perturbations in physiology and morphology can impact the subsequent juvenile and
adult phenotypes in these animals (see Chapter 2 for discussion). Thus it is difficult to
interpret whether the variations in physiology of chicken embryos cultured on a variety
of foreign yolks mirror the attributes that are characteristic of the species which
represents the yolk.
In other words, does the species-specific variation in yolk
environment allow the chicken embryo to “adopt” the physiological characteristics of the
species from which the yolk environment was derived? The answer to this question
must await further study of early embryonic physiology of birds on a comparative scale.
It is important to consider that with respect to yolk TE, initial egg mass, and
incubation period, the inclusion of the Ratitae in the regression models significantly
70
altered the nature of the relationship between these factors and the measured
embryonic traits by either rendering the model non-significant or drastically changing the
slope of the regression (Fig, 3.5B,C,D, Fig. 3.6B,C, D, and Fig. 3.7B, C). In contrast,
yolk T3 appeared to predict body mass and heart rate in a similar fashion, regardless of
the inclusion or exclusion of Ratitae (Fig. 3.5A and 3.6A). These findings highlight the
need to take phylogenetic taxa into consideration when attempting to generate models
that are designed to predict physiological and morphological attributes of developing
animals.
Summary
In summary, early avian embryonic traits such as heart rate, metabolic rate and
body mass can be significantly changed by altering the yolk environment in a speciesspecific manner. The underlying mechanism of this phenomenon is correlated with the
inter-species variation in yolk hormone concentrations, T 3 and TE, and other potential
yolk factors that scale with egg mass and incubation period of the egg.
The low
coefficient of determination of the regression models indicated that the factors included
in this study are not the only factors to contribute to inter-species variation in early
embryonic phenotype. Further identification of potential yolk factors will be fruitful when
testing the exciting possibility that differences in yolk constituents that affect
developmental patterns during embryonic stages of life influence the phenotypes of latestage embryos, hatchlings, juveniles and adults of avian species.
In Chapter 4, I further elucidate the role of yolk environment in the determination
of early embryonic phenotypes of two chicken breeds, the layer and the broiler, which
71
show largely different growth rates during juvenile development and marked differences
in adult muscle structure and body mass.
72
Figure 3.1 Schematic of embryo culture method. Panel A represents the preparation of yolk culture dishes. Panel B
represents the removal of embryos from original yolk via a filter paper ring, and the placement onto prepared yolk dishes.
73
Table 3.1 Egg components and length of prenatal development (incubation period) of birds. Values presented as
mean + S.E.M.
Parvclass, Order, Species, Breed
Incubation
Period
(range)
T3
n
Galloanserae
Galliformes
Quail
Chicken
White Leghorn
Cornish Rock Broiler
Buff Cochin
Buff Cochin Bantam
White Leghorn Bantam
Narragansett Turkey
Anseriformes
Duck
Blue Indian Runner
Khaki Campbell
Mallard
Buff Runner
Chinese Goose
Ratitae
Struthioformes
Emu
Ostrich
16.5
(14 - 19)
21
21
21
20
(19 – 21)
20
(19 – 21)
28
28
28
28
(26 – 30)
28
30
(29 – 21)
56
(50 -62)
41
(40 – 42)
TE
41
23
3
Egg Mass
(g)
66.83 + 5.59
45.99 + 9.17
14.04 + 0.56
Yolk Conc.
(pg mg-1)
1.55 + 0.13
1.01 + 0.11
1.24 + 0.13
17
4
3
3
4
47.44 + 3.81
59.53 + 1.37
48.80 + 3.35
66.92 + 2.03
35.62 + 1.14
3
119
85
19
Egg Mass
(g)
54.20 + 2.94
41.18 + 2.39
11.90 + 3.35
Yolk Conc.
(pg mg-1)
4.24 + 0.20
4.10 + 0.25
3.79 + 0.56
0.89 + 0.27
1.05 + 0.10
0.46 + 0.09
1.20 + 0.53
0.59 + 0.15
60
12
12
12
12
47.90 + 1.95
60.79 + 1.52
51.669 + 1.08
66.19 + 1.07
36.08 + 0.51
4.13 + 0.30
3.32 + 0.55
4.63 + 0.58
4.88 + 0.84
3.59 + 0.69
30.75 + 0.67
1.13 + 0.14
12
30.14 + 0.67
4.23 + 0.66
3
18
14
3
4
3
70.70 + 4.72
91.14 + 7.77
74.31 + 3.45
71.417 +3.65
74.77 + 3.39
77.96 + 3.27
1.42 + 0.29
2.19 + 0.17
2.37 + 0.41
2.39 + 0.33
1.82 + 0.23
3.16 + 0.40
6
34
28
6
4
6
72.28 + 3.13
84.83 + 5.149
71.45 + 1.20
69.48 + 2.42
74.77 + 3.39
74.18 + 2.99
4.82 + 0.73
4.58 + 0.33
4.36 + 0.31
3.99 + 0.45
3.52 + 0.48
3.09 + 0.68
4
4
73.228 + 3.05
150.06 +3.65
2.30 + 0.33
1.56 + 0.13
12
6
69.95 + 1.70
147.25 + 2.935
5.46 + 0.39
5.62 + 1.18
6
6
3
1145.73 + 597.49
1145.73 + 597.49
630.96 + 25.718
1.33 + 0.16
1.33 + 0.16
1.63 + 0.15
17
17
13
794.58 + 108.67
794.58 + 108.67
594.75 + 15.75
2.39 + 0.21
2.39 + 0.21
2.44 + 0.23
3
1660.50 + 66.90
1.03 + 0.10
4
1660.50 + 66.90
2.28 + 0.51
74
n
1.9
1.8
IP(Neornithes) = 0.89 IEM0.28(0.01)
2
log IP (days)
1.7
r = 0.86, n =130, p < 0.001
1.6
1.5
1.4
1.3
1.2
IP(Galloanserae) = 0.94 IEM0.25(0.01)
2
r = 0.76, n =114, p < 0.001
IP(Galliformes) = 1.03 IEM0.18(0.01)
2
r = 0.75, n = 80, p < 0.001
0.0
0.0 1.0 1.2 1.4 1.6 1.8 2.0 2.2 2.4 2.6 2.8 3.0 3.2 3.4 3.6
log IEM (g)
FIGURE 3.2 Relationship between initial egg mass (IEM) and incubation period (IP) for
7 species of birds (see Table 3.1). Regression plots indicate IEM-IP relationships
among Neornithes (black symbol, solid line), Galloanserae (grey symbol, long-dashed
line) and Galliformes (white symbol, short-dashed line). Least-squares linear
regressions, α < 0.05.
75
Table 3.2 Least-squares regression estimation on log 10-transformed data for the relationship between yolk hormone
concentrations (yolk T3 or yolk TE) and IEM.
yolk T3 = aI EMb
yolk TE = aI EMb
n
a
b (SE)
r2
p
n
a
b (SE)
r2
p
46
-0.036
0.080 (0.076)
0.023
0.302
129
0.659
-0.061(0.041)
0.017
0.137
Galloanserae
43
-0.597
0.409 (0.155)
0.145
0.012
114
0.265
0.190(0.067)
0.066
0.006
Ratitae
6
1.461
0.452(0.107)
0.816
0.014
15
0.870
-0.184(0.183)
0.072
0.333
Neornithes
Table 3.3 Least-squares regression estimation on log10-transformed data for the relationship between IP and yolk
hormone concentrations (yolk T3 or yolk TE).
IP = a yolk T3b
IP = a yolk TEb
n
a
b (SE)
r2
p
n
a
b (SE)
r2
p
49
1.392
0.168(0.070)
0.109
0.020
135
1.429
-0.077(0.054)
0.015
0.158
Galloanserae
43
1.354
0.163(0.040)
0.283
<0.001
119
1.289
0.097(0.034)
0.067
0.005
Ratitae
6
1.625
0.508(0.146)
0.751
0.026
15
0.475
-0.055(0.033)
0.0675
0.350
Neornithes
76
A
0.8
log yolk T3 (pg mg-1)
0.6
yolk T3 = -0.597 IEM
0.41(0.15)
2
0.4
r = 0.145, n = 43, p = 0.012
0.2
0.0
-0.2
-0.4
yolk T3 = 1.461 IEM
0.45(0.11)
2
r = 0.816, n = 6, p = 0.014
-0.6
-0.8
0.0 1.0 1.2 1.4 1.6 1.8 2.0 2.2 2.4 2.6 2.8 3.0 3.2 3.4
B
1.2
log yolk TE (pg mg-1)
1.0
0.19(0.07)
yolk TE = 0.265 IEM
2
r = 0.066, n = 114, p = 0.006
0.8
0.6
0.4
0.2
0.0
0.0 1.0 1.2 1.4 1.6 1.8 2.0 2.2 2.4 2.6 2.8 3.0 3.2 3.4
log IEM (g)
FIGURE 3.3 Relationship between log initial egg mass (IEM) and log yolk T 3 (A) or log
yolk TE (B) for 7 species of birds (see Table 3.1). Significant regression plots indicate
relationships among Galloanserae (white symbol, solid line) and Ratitae (black symbol,
solid line).
77
A
1.8
1.7
log IP (days)
1.6
IP = 1.39 yolk T30.17(0.07)
1.5
2
r = 0.11, n = 49, p = 0.02
1.4
1.3
1.2
IP = 1.35 yolk T 0.16(0.04)
3
2
r = 0.28, n =43, p < 0.001
0.0
-0.8
-0.6
-0.4
-0.2
0.0
0.2
0.4
0.6
0.8
-1
log T3 (pg mg )
B
1.8
1.7
log IP (days)
1.6
NS
1.5
1.4
1.3
1.2
IP = 1.29 yolk TE0.10(0.03)
2
r = 0.07, n = 135, p = 0.005
0.0
0.0
0.2
0.4
0.6
0.8
1.0
1.2
-1
log yolk TE (pg mg )
FIGURE 3.4 Relationship between log yolk T3(A) or log yolk TE (B) and log IP for 7
species of birds (see Table 3.1). Significant regression plots indicate relationships
among Neornithes (broken line) and Galloanserae (solid line). Black symbols represent
Ratitae removed from the model. Non-significant regression denoted by NS.
78
Table 3.4. Morphological and physiological traits of White Leghorn chicken (Gallus gallus) cultured on yolks of different
bird species. Values presented as mean + S.E.M
Parvclass, order, species,
breed
Neognathae
Galliformes
Quail
WL Chicken
Narragansett Turkey
Anseriformes
Duck
Blue Indian Runner
Khaki Campbell
Mallard
Buff Runner
Chinese Goose
Palaeognathae
Struthioformes
Emu
Ostrich
Wet BM
fH
Mass-specific
HH stage
(bpm)
175 + 1
173 + 2
164 + 3
172 + 3
192 + 3
180 + 2
180 + 3
188 + 3
163 + 5
192 + 5
175 + 2
179 + 3
n
77
51
19
21
11
26
11
(ml g-1 h-1)
1.98 + 0.24, 77
3.54 + 0.30, 51
4.56 + 0.48, 19
4.14 + 0.42, 21
1.80 + 0.54, 11
1.98 + 0.42, 26
4.56 + 0.72, 11
n
85
54
21
22
11
31
11
17.5 + 0.1
17.4 +0.1
17.3 + 0.6
17.3 + 0.2
18.0 + 0.2
17.7 + 0.1
17.6 + 0.2
11
4.56 + 0.72, 11
11
17.6 + 0.2
14.64 + 1.15
n
291
198
50
118
30
93
52
9
18
22
3
41
15
3.12 + 0.36, 15
20
17.7 + 0.2
11.50 + 0.76
12.50 + 0.89
8.63 + 0.87
103
64
39
177 + 3
176 + 3
180 + 5
29
21
8
4.38 + 0.36, 29
4.50 + 0.48, 21
4.14 + 0.30, 8
31
20
9
17.7 + 0.1
17.6 + 0.2
17.8 + 0.3
n
83
52
20
21
11
31
11
(mg)
12.18 +0.49
44 + 0.55
11.12 + 1.02
10.31 + 0.70
14.16 + 0.92
13.43 + 0.91
11.21 + 1.30
11
11.21 + 1.30
20
31
23
8
79
1.6
1.6
A
BM = 1.01 yolk T3
2
r = 0.056, n = 83, p = 0.03
BM = 0.87 yolk TE0.33(0.11)
r2 = 0.073, n = 114, p = 0.004
1.4
log BM (mg)
log BM (mg)
1.4
B
0.40(0.18)
1.2
1.0
1.2
1.0
0.8
0.8
BM = 0.63 yolk TE0.70(0.18)
r2 = 0.16, n = 83, p < 0.001
BM = 1.00 yolk T30.35(0.17)
2
r = 0.056, n = 114, p = 0.04
0.0
-0.05
0.00
0.05
0.10
0.15
0.20
0.0
0.25
0.0
0.30
0.4
0.5
0.6
1.6
1.6
C
1.4
D
1.4
NS
log BM (mg)
NS
log BM (mg)
0.8
log yolk TE (pg mg-1)
log yolk T3 (pg mg-1)
1.2
1.0
0.8
1.2
1.0
0.8
0.10(0.04)
0.0
0.7
BM = 0.39 IP0.49(0.16)
r2 = 0.10, n = 83, p = 0.003
BM = 0.88 IEM
r2 = 0.071, n = 83, p = 0.01
0.0
0.0
1.0
1.5
2.0
2.5
3.0
3.5
0.0
log IEM (g)
1.2
1.4
1.6
log IP (days)
80
1.8
FIGURE 3.5 Relationship between log body mass (BM) and log yolk T 3(A), log yolk TE (B), log IEM(C), or log IP(D) for
chicken embryos cultured on the yolks of 7 species of birds (see Table 3.1). Regression plots indicate relationships
among Neornithes (broken line, white and black symbol) and Galloanserae (solid line, white symbols). Black symbols
represent Ratitae.
81
2.5
2.5
A
fH(Galloanserae) = 2.23 yolk T30.09(0.03)
NS
2.4
log fH (bpm)
2.4
log fH (bpm)
B
r2 = 0.04, n = 291, P < 0.001
2.3
2.2
2.1
2.3
2.2
2.1
fH = 2.18 yolk TE0.11(0.04)
fH(Neornithes) = 2.23 yolk T30.08(0.03)
r2 = 0.02, n = 291, P = 0.02
r2 = 0.03, n = 394, P < 0.001
0.0
-0.1
0.0
0.0
0.1
0.2
0.3
0.4
0.5
0.0
0.4
-1
2.5
2.5
fH = 2.17 IEM
log fH (bpm)
log fH (bpm)
2.2
fH = 2.00 IP0.19(0.04)
2.3
2.2
fH = 2.17 IP0.05(0.02)
r2 = 0.01, n = 394, P = 0.02
0.0
r2 = 0.01, n = 394, P = 0.02
0.0
1.0
1.5
2.0
2.5
0.8
2.1
fH = 2.21 IEM0.13(0.005)
0.0
0.7
r2 = 0.07, n = 291, P < 0.001
2.4
2.3
2.1
D
0.04(0.01)
r2 = 0.04, n = 291, P < 0.001
2.4
0.6
log yolk TE (pg mg-1)
log yolk T3 (pg mg )
C
0.5
3.0
3.5
0.0
log IEM (g)
1.2
1.4
1.6
log IP (days)
82
1.8
FIGURE 3.6 Relationship between log heart rate (fH) and log yolk T3(A), log yolk TE (B), log IEM(C), or log IP(D) for
chicken embryos cultured on the yolks of 7 species of birds (see Table 3.1). Regression plots indicate relationships
among Neornithes (broken line, white and black symbol) and Galloanserae (solid line, white symbols). Black symbols
represent Ratitae.
83
.
1.2
A
1.0
log mass-specific VO2 (ml g-1 h-1)
log mass-specific VO2 (ml g-1 h-1)
1.2
NS
0.8
.
0.6
0.4
NS
0.2
0.00
0.05
0.10
0.15
0.20
0.25
r2 = 0.07, n = 77, P = 0.24
0.8
0.6
0.4
.
0.2
VO2 = -1.01 yolk TE-0.36(0.15)
r2 = 0.02, n = 106, P = 0.02
0.0
0.30
VO2 = -0.86 yolk TE-0.60(0.26)
1.0
0.0
0.0
-0.05
.
B
0.4
1.0
NS
0.8
.
0.6
0.4
0.2
0.0
.
VO2 = -1.03 IEM-0.18(0.06)
r2 = 0.05, n = 77, P = 0.04
0.0
0.7
0.8
1.2
C
log mass-specific VO2 (ml g-1 h-1)
log mass-specific VO2 (ml g-1 h-1)
.
0.6
log yolk TE (pg mg-1)
log yolk T3 (pg mg-1)
1.2
0.5
1.0
1.5
D
1.0
NS
0.8
0.6
0.4
0.2
NS
0.0
2.0
2.5
3.0
0.0
3.5
1.2
1.4
1.6
log IP (days)
log IEM (g)
84
1.8
FIGURE 3.7 Relationship between log mass-specific oxygen consumption rate ( ) and log yolk T3(A), log yolk TE (B),
log IEM(C), or log IP(D) for chicken embryos cultured on the yolks of 7 species of birds (see Table 3.1). Regression plots
indicate relationships among Neornithes (broken line, white and black symbol) and Galloanserae (solid line, white
symbols). Black symbols represent Ratitae.
85
CHAPTER 4
INFLUENCE OF INTER-BREED YOLK FACTORS ON HEART RATE, OXYGEN
CONSUMPTION, DEVELOPMENT RATE AND BODY MASS OF EARY-STAGE
CHICKEN EMBRYOS
Introduction
The layer chicken and the broiler chicken significantly differ in embryonic,
juvenile and adult phenotype. Layers are predominantly smaller in size and are prolific
egg-layers, while broilers reach maturity in a short period of time and attain larger adult
mass for meat-yield. Broilers are often four times the size of layers at 42 days posthatch, and have significantly different muscle morphology and physiology due to their
fast growth rates (McRae et al., 2006). As a result of extremely fast muscle tissue
growth, broilers suffer from ascites, a syndrome that occurs when the oxygen demands
of muscle tissue exceeds the capabilities of the cardiovascular system to deliver oxygen
to these tissues (Olkowski, 2007). This syndrome has been equated to pulmonary
hypertension in humans (Martinez-Lemus et al., 2000).
The differences in morphological and physiological traits between mature layer
and broiler chickens can be traced back to ontogenic differences during embryogenesis
(Latimer and Brisbin, 1987; Burggren et al., 1994; Clum et al., 1995; Mitchell and Burke,
1995; Schreurs et al., 1995; Martinez-Lemus et al., 1998; Martinez-Lemus et al., 1999;
Koenen et al., 2002; Odom et al., 2004). Embryonic mass differ significantly between
layers and broilers throughout incubation, with differences becoming more apparent
towards the end of incubation (Pal et al., 2002). Also, broiler embryos have a faster rate
of development than layer embryos beginning at the first 50 hours of development
86
(Boerjan, 2005).
In fact, during the last 5 days of incubation, broiler embryonic
metabolic heat production becomes significantly greater than layer embryonic metabolic
heat production due to the larger body mass of broilers (Sato et al., 2006). When body
mass difference between the two breeds was accounted for, layer heat production per
gram of body weight was significantly greater than broiler heat production per gram of
body weight (Sato et al., 2006). Heart morphology also differs between the two breeds;
with only broiler embryos showing ventricular hypertrophy as early as 40 hours of
incubation (Boerjan, 2005). Additionally, heart rates of broiler embryos are higher than
that of layer embryos, with this difference significantly reversing during the first two days
after hatching (Yoneta et al., 2006; Yoneta et al., 2007).
Given the genetic differences between broilers and layers, it is generally
assumed that direct genetic effects are responsible for these observed phenotypic
difference between the two breeds (Dunnington, 1994; International Chicken
Polymorphism Map Consortium, 2004).
However, the difference in genotype may
indicate a difference in maternal condition which may also give rise to indirect genetic
effects (i.e. maternal effects) in the subsequent generation.
This conjecture is
supported by evidence that egg yolk hormones of maternal origin such as testosterone
(TE) and triiodothyronine (T 3) vary significantly among and within avian species, and
have significant and permanent effects on offspring phenotype (Wilson and McNabb,
1997; Groothuis et al., 2005).
Currently, the role of epigenetic processes such as
maternal effects in the determination of broiler and layer phenotypes remains largely
unexplored.
87
I hypothesized that breed-specific embryonic phenotypes can be attributed to the
actions of breed-specific, maternally-derived yolk factors.
To test this hypothesis, I
utilized a novel, ex ovo method which allowed complete alteration of the yolk
environment of intact chicken embryos during a 48 hour period in development, starting
as early as Hamburger and Hamilton stage 9 (HH9; 1950).
development rate, heart rate, and
The body mass,
of layer and broiler embryos that developed within
the context of breed-specific alteration in yolk environment were investigated.
Subsequently, I quantified levels of T 3 and TE in the yolks of broiler and layer eggs to
determine if these physiologically-relevant egg factors contribute to the growth and
cardiac chronotropic differences observed between broilers and layers.
Materials and Methods
Incubation
White Leghorn layer eggs and Cornish Rocks broiler eggs were purchased from
Texas A&M University (College Station), Ideal Poultry, Breeding Farms, Inc. (Cameron,
TX), and Moyer‟s Chicks (Quakertown, PA). Upon arrival, eggs were stored at room
temperature (23-26°C) until the start of incubation.
commercial incubators at 37°C and 58% humidity.
All eggs were incubated in
Avian eggs were automatically
turned every hour during the course of the incubation period. Proper humidity and
temperature were checked daily.
Triiodo-L-thyronine Radioimmunoassay
Fresh, unincubated broiler and layer chicken eggs were dissected, and yolks
were separated, homogenized and stored at -20 °C for assays.
The methanol/
chloroform extraction procedure of Wilson and McNabb (1997) was slightly modified to
88
increase T3 extraction efficiency. 0.5 g of previously homogenized yolk was placed in
13 ml glass conical tube and 2 ml of methanol (supplemented with 1mM propylthiouracil
(PTU)) was added. The components were mixed (via vortex mixer) for approximately 5
minutes to ensure a homogenous suspension. Labeled thyroid hormone was added to
each sample (~2000 cpm of
125
I-T4 or
125
and counted using a gamma counter.
I-T3; MP Biomedicals, New York, NY), mixed
Subsequently samples were shaken (150
oscillations/min on a shaker) for 10 min., and centrifuged for 10 minutes at 2700 rpm.
The supernatant was decanted into a 13ml glass conical tube and the precipitate was
resuspended in 1ml methanol (1mM PTU) and shaken, spun and decanted (as above)
in another 13ml glass conical tube. Five ml of chloroform and 0.5 ml of 2N ammonium
hydroxide was added to each tube, and the tubes were shaken and centrifuged. The
upper phase of corresponding tubes were removed and combined in a 13ml glass tube.
The suspension was allowed to dry overnight under a filtered air stream. The dried
samples were resuspended in 1 ml 2N ammonium hydroxide, shaken, centrifuged and
decanted into a 13ml tube. One ml of chloroform was added to each tube, and the
tubes were shaken, centrifuged, and the upper phase was removed and allowed to dry
under a filtered air stream.
The samples were resuspended in 150 µl phosphate
buffered saline (supplemented with alizarin) and counted (1 min).
The extraction
efficiency was calculated for each sample by calculating the percentage of radioactivity
that remained after the extraction procedure.
T3 concentrations of the resuspended sample (from the last step of extraction
above) were analyzed using competitive binding radioimmunoassay kits (Free T 3
89
Antibody Coated Tube-125I RIA Kit, MP Biomedicals, New York, NY). Procedures of
the hormone kit were followed exactly.
Testosterone Radioimmunoassay
Testosterone concentrations in the previously collected yolk samples were
determined via radioimmunoassay following the protocol for steroid hormones in egg
yolks established by Schwabl (1993) and modified for use with broiler and layer chicken
yolks. The protocol was modified by using a phosphate buffered saline containing yolk
stripped of hormones as the background buffer for the standard curves. Using yolk
pooled from eggs collected from all species controlled for extra yolk material present in
samples via direct assays.
This negated the need for the use of column
chromatography to clean yolk samples. The protocol for stripping hormones from yolk
samples was modified from that of Wingfield et al. (1984) for stripping plasma of
steroids.
Egg Components Mass and Egg Mass
Egg components mass expressed in g include fresh yolk mass, fresh albumen
mass and fresh shell mass. Total egg mass is the sum of these components. Masses
were obtained by first weighing the intact egg using a balance (Denver Instruments
Company, CO), and then subsequently separating and prepping yolk and shell
components for weighing. Yolks were individually placed in plastic dishes and cleaned
of residual albumen using Kimwipes. Shells were washed of residual albumen and
allowed to dry for at least 24 h prior to weighing. Since significant amounts of albumen
were permanently lost in the process of separating egg components, albumen mass
was obtained by subtracting shell and yolk mass from total egg mass.
90
Embryo Cultures
Layer and broiler embryos (~40 hours of incubation; Hamburger and Hamilton
(1951) stages 9-14) were removed from their original yolks and transferred to parafilm
culture dishes filled with the yolk of a different chicken breed by following the methods
previously outlined by Chapman et al. (2001). Parafilm culture dishes were prepared as
previously described by Chapman et al. (2001), with the exception of the culture
medium used. Parafilm culture dishes were prepared by covering the lids of 35mm
petri-dishes with parafilm, and completely filling them with Yolk-Tyrode (YT) medium
through a square opening (2 cm x 2cm) cut out of the parafilm (see Chapter 3, Fig.3.1).
YT medium consisted of one part fresh, homogenized yolk from layer or broiler eggs,
and one part 1X Tyrode‟s Solution supplemented with 5 units penicillin-streptomycin/ml
(v/v; Sigma T1788, St. Louis, MO).
Embryos (HH 9-14) and associated vitelline membranes were carefully excised
from their native yolk using Whatman filter paper rings as the carrier. Embryo removal
from the yolk sac consisted of first removing albumen from the area surrounding the
embryo. Then a filter paper ring (outer diameter = 38 mm; inner diameter = 15 mm)
was positioned on the vitelline membranes so that the embryo was centrally framed by
the ring, and the surrounding membranes tightly adhered to the filter paper ring. The
membrane along the outer edge of the filter paper ring was cut and the paper ring
carrying the embryo was lifted from the yolk using fine-tipped forceps. Embryos were
washed free of residual yolk and albumen by gently swirling in three washes of warm 1X
Tyrode‟s, penicillin/streptomycin. The edge of the filter paper ring carrying the embryo
was blotted on a Kimwipe for a few seconds to remove excess liquid from the embryo,
91
and a second filter paper ring (outer diameter = 38 mm, inner diameter = 30 mm) was
placed on top of the embryo/filter paper as to sandwich the embryo and associated
vitelline membranes between the two filter paper rings (Fig. 3.1).
This entire
embryo/filter paper complex was then placed onto prepared parafilm culture dishes, with
the embryo dorsal side down, and in direct contact with the YT medium at the opening
in the parafilm dish.
Seven parafilm embryo cultures were placed into a large petri dish (135 mm X
10mm), which then was placed into a seal-top plastic chamber (170 mm diameter, 80
mm height) outfitted with inflow and outflow ports on the lid. Two Kimwipes, wetted with
10 ml of distilled water, were placed at the bottom of the plastic chamber to maintain a
high level of humidity. After the lids were placed on the containers, the containers were
flushed via the ports for 20 minutes with a gas mixture of 95% oxygen (O 2) and 5%
carbon dioxide (CO2; Wösthoff Gas Mixer, Calibrated Instruments, Inc., NY).
After
flushing of chambers, ports were sealed, and the whole container was maintained in a
37.5 + 0.2°C, clear-top Styrofoam incubator (Circulated Air Hova-Bator Incubator, Model
No. 1590, GA). The start of experiments was considered to be 1.5 hours after the
chambers have been sealed and placed in incubators.
The high oxygen atmosphere (95% O 2, 5% CO2) provided for the cultures in this
study was to increase survivability and proper development of cultured embryos.
In
vitro cell cultures are commonly maintained under this oxygen to carbon dioxide
pressure to prevent acidification of culture medium (personal communication with Susan
Chapman). Heart rate of culture embryos incubated under this atmosphere was similar
to that of windowed embryos (Chapter 2), and embryonic development progressed
92
normally according to Hamburger and Hamilton (1951). The actual percent oxygen in
the chambers declined over the period of parafilm embryo culture incubation. Initial
reading was 90+5% O2, and decreased to 47+8% O2 after 24 h of incubation in a cohort
of 5 chambers tested.
For each trial, variability in the rate of PO 2 decline among
chambers was accounted for by evenly distributing experimental groups among all
chambers.
Wet and Dry Body Mass
Live embryos were anesthetized via exposure to -20°C for 4 min. Fresh embryo
mass was determined by completely excising the embryo from the surrounding vitelline
membranes in 1X Tyrode‟s solution.
Embryos were then carefully blotted with
Whatmann filter paper for 20 seconds to remove all visible water surrounding the
embryo, and weighed using a microbalance. Dry embryo mass was obtained placing
the fresh embryos into a 60°C oven for 48 h. Embryo wet mass was assessed at 24h
and 48h after the start of the experiment, and dry weight was assessed at the end of
48h only.
Development Rate
Embryonic development rate was determined by staging embryos according to
Hamburger and Hamilton (HH) stages at the time of parafilm embryo culture and then
again at 12, 24 and 48 h after transfer. Before embryos were placed on parafilm culture
dishes, they were placed in a petri-dish filled with warm 1X Tyrode‟s and viewed at 10X
magnification under a light microscope for staging of embryos (HH9-HH14) by counting
somite numbers. Staging after 24 and 48 h of incubation was primarily based on the
degree of spine curvature (HH14 and older) since embryos could not be magnified to
93
assess somite number or limb development. For example, HH14 was indicated by the
forebrain and hindbrain forming a 45° angle, while HH15 was indicated by 67.5°
curvature of the forebrain and hindbrain.
Broiler and layer embryos assigned to
treatment groups with respect to yolk type, were equally distributed between yolk
groups according to HH stage.
Heart Rate
The heart beat of embryos maintained in clear-top incubator (Circulated Air
Hova-Bator Incubator, Model No. 1590, GA) set at 37.5+°C was visually counted over
the course of one min.
Heart rates (fH) were reported as beats per minute (bpm).
Heart rate was recorded at 12, 24 and 48 h after the start of the experiments.
Mass-Specific Metabolic Rate
Oxygen consumption of embryos was assessed by closed-system respirometry.
Individual metabolic respirometers were constructed from 118 ml mason jars with metal
screw-top lids. The bottom of the respirometer was filled with polyester casting resin to
reduce the respirometer air volume to 27 ml. The lid was outfitted with a syringe port
constructed from the hub of a 25 gauge needle to allow for the attachment of a metal
stopcock and a 5ml glass syringe (Proper MFG Co., Inc.) for air sampling. Individual
embryo cultures were placed in the center of each respirometer, and the lids were
tightly sealed to the mouth of the respirometer with the aid of petroleum jelly. The
repirometers were then placed into an acrylic reptile incubator set at at 37.5°C. Each
embryo in its respirometer was allowed to equilibrate for 30 min with the syringe port
open for free exchange of air between the respriometer and the ambient air.
94
Thirty min after placement of respirometers into the reptile incubator, a small 3way metal stopcock attached to a 5ml-glass syringe was inserted into the syringe port of
the lid, and a 1.5 ml sample of air from the respirometer was drawn into the syringe and
injected into a flow-through oxygen electrode (Model 16-730 Microelectrodes, Inc., NH)
to obtain the initial PO2 of gas in the respirometer. The voltage output of the electrode
was converted into partial pressure by Chart software (Power Lab:
Chart 5,
ADInstruments). The stopcock-syringe was returned to the port of the respirometers,
and the respirometers were sealed by setting the stopcock to allow passage of air
between the respirometer and the syringe only. After 60 min, 1.5 ml of respirometer gas
was drawn into the syringe, and the gas in the syringe was injected into the flow-through
the oxygen electrode to obtain the final PO2 value (as described above). The massspecific oxygen consumption rate (
in mlO2.g.h-1) of transfer embryos was
calculated using the following equation:
Where ∆PO2 is the change in oxygen pressure of the gas over time interval (mmHg), v is
total volume of air in the respirometer (ml), Δt is the time elapsed between PO2 readings
(h), and m is the wet mass (g) of the embryo. Total gas volume was determined for
each metabolic respirometer by subtracting the weight of the empty respirometer from
the weight of the chamber filled with distilled water. An additional 5 g was subtracted
from the final weight to account for the volume taken up by the parafilm culture dish,
assuming that 1 g of distilled water occupied a volume of 1 ml.
Respirometers
containing yolk culture dishes without embryos were sampled during each trial run to
95
account for non-biological fluctuations in oxygen concentration of respirometer gas.
Mass-specific
of embryos was adjusted accordingly.
Oxygen consumption was assessed at 12, 24 and 48 hours after the start of the
experiment in order to capture a full range of developmental stages and body masses
for each experimental group.
Statistical Analyses
Changes in wet body mass, heart rate and yolk hormone concentrations were
assessed via either factorial three-way or two-way ANOVAs with respect to embryo
type, yolk type and time, depending on the number of contributing factors, followed by
Tukey‟s post-hoc comparisons. Since the design of the ANOVA analyses did not permit
post-hoc comparison between embryos cultured on their native yolks (broiler embryos
cultured on broiler yolk and layer embryos cultured on layer yolk) additional, two-way
ANOVAs with respect to group (defined by embryo type and yolk type) and time were
performed. HH stage data were considered to be non-discrete, so non-parametric twoway ANOVAs and subsequent Kruskall-Wallis post-hocs were used to detect statistical
significance among experimental groups.
Yolk hormone concentrations were
normalized via logarithmic transformations prior to statistical analyses.
Data were
transformed back for graphical presentation.
The relationship between egg component mass (albumen, yolk or shell) and egg
mass was assessed using regression analyses.
The allometry of egg components
mass to egg mass was investigated by regression of logarithm of each egg
component‟s mass on the logarithm of egg mass (Dzialowski and Sotherland, 2004).
The slope (b) of the regression line was used to determine if the relationship was
96
isometric (b = 1) or allometric (b < 1 or b > 1). Regressions were considered isometric
when the 95% confidence intervals of the slope includes 1, and allometric when it does
not include 1. Isometry indicates a linear relationship between egg components mass
and egg mass, while allometry indicates non-linear relationship between the two
variables. Slope data reported as b + 95% confidence interval.
Results
Body Mass
Wet and dry mass of embryos are depicted in Figures 4.1A and B, respectively.
As anticipated, embryo wet mass significantly increased from ~12.70 + 5.10g to ~22.29
+ 7.75 g over the course of 48 hours of development (F (1,
131)
= 59.06, p < 0.001).
Interestingly, during this early development (~40 to 88 h of age), layer embryos cultured
on broiler yolk for 24 and 48 h were significantly heavier, in terms of wet mass, than
layer embryos cultured on layer yolk. However, they were not different from broiler
embryos cultured on broiler yolk (Tukey‟s, q = 3.0, p = 0.040, and Tukey‟s, q = 3.08, p =
0.03, respectively). After 48 h of culturing, dry mass of layer embryos showed the
same significant trends as wet mass, with layers cultured on broiler yolk showing
heavier mass than those cultured on layer yolk (Tukey‟s, q = 3.38, p = 0.03). However,
yolk type did not significantly alter broiler embryo wet or dry mass at any time during the
course of the experiment. It appeared that a change in yolk environment affected layer
and broiler embryos differentially, with layer embryos showing significant alterations in
breed-specific morphology in response broiler yolk environment.
97
Egg Component Masses
For broiler and layer breeds, albumen, yolk and shell mass were directly related
to egg mass as depicted in Figure 4.6.
Although the relationship between egg
component mass and egg mass (represented by slope of regression lines, b) do not
significantly differ between the two breeds for each egg component mass, the
elevations (y-intercept) of the regressions do in fact significantly differ between broilers
and layers for each egg component. Over the range of egg mass, broiler albumen
mass and shell mass were significantly lower than layer albumen mass and shell mass
(Fa(1, 87) = 31.73, pa < 0.0001; Fs(1, 87) = 7.33, ps = 0.008), while yolk mass of broiler eggs
was greater than that of layer eggs (F y(1, 87) = 75.33, py < 0.0001). However, egg mass
was not different between broiler and layer eggs (58.47+0.39 and 58.62+1.15 g,
respectively; t = 0.15, p = 0.89).
The comparison of the slopes (b) of the log-log regressions between each egg
component‟s mass and egg mass indicated that the relative contribution of yolk mass to
egg mass changed in direct proportion to initial egg mass of broiler eggs (b = 0.85 +
0.16, r2 = 0.48) and layer eggs (b = 0.85 + 0.16, r2 = 0.33). However, the relative
contribution of albumen mass and shell mass to egg mass increased allometrically with
egg mass for both broiler (ba = 1.32 + 0.10, ra2 = 0.85; bs = 0.67 + 0.15, rs2 = 0.39) and
layer eggs (ba = 0.67 + 0.15, ra2 = 0.39; bs = 0.73 + 0.14, rs2 = 0.32).
Development Rate
Over the course of the experiment, embryos of all experimental groups showed
significantly more mature developmental stages from one time period to the next
(Tukey‟s, p‟s < 0.0001; Fig. 4.2). A comparison of HH stage between broiler embryos
98
and layer embryos at the start of the experiment (40 h old embryos, in ovo), revealed
that broiler embryos were significantly less mature than layer embryos (Tukey‟s, q =
2.84, p = 0.044). The only difference detected among the four culture groups at 0 h was
between layer embryos cultured on broiler yolk and broiler embryos cultured on broiler
yolk (LB > BB; Tukey‟s, q = 2.97, p = 0.04). By 12 h after the start of incubation on the
parafilm culture dishes, layer and broiler embryos no longer showed any significant
difference in HH stage, and there were no differences among any culture groups.
However, after 24 h had elapsed from the start of the experiment, yolk type (F(1, 148) =
5.39, p = 0.022), but not embryo type had a significant effect on embryo stage. At this
time, layer embryos residing on layer yolk displayed less mature HH stage than did
layer embryos residing on broiler yolk (Tukey‟s, q = 3.40, p = 0.016). After 48 h from
the start of the experiment, layer embryos were more developmentally advanced than
broiler embryos (F(1, 77) = 4.15, p = 0.045), but post-hoc comparisons among groups did
not reveal any significant differences between groups. Similar to the effects observed of
body mass among culture groups, breed-specific change in yolk environment
significantly altered the development rate of layer embryos, but not that of broiler
embryos.
Mass-Specific Oxygen Consumption
For this study, mass-specific
of cultured embryos ranged from approximately
5 to 24 ml g-1 h-1 for embryo masses of 4 to 24 g. (Fig. 4.3A). The relationship between
mass-specific metabolic rate (ml g-1 h-1) and wet mass (g) of each experimental group
(defined by embryo type (B or L) and yolk type (B or L)) was characterized in Figure
4.3B.
Comparisons of the regression models revealed that neither the slopes nor
99
elevations of the regression lines were significantly different from one another (F (3, 34) =
0.85, p = 0.48; F(3,
37)
= 0.53, p = 0.66, respectively).
Thus, for this study, the
relationship between mass-specific metabolic rate and wet mass of cultured chicken
embryos (ranging from 3.3 to 23 g) can be expressed as
= -0.75 M + 18.79 (F =
62.184, r2 = 0.61, p < 0.001; Fig. 4.4A).
Heart Rate
Comparison of mean fH of broiler and layer embryos in their native yolk
environments (broilers cultured on broiler yolk, and layers cultured on layer yolk) over
the 36 hour period after the start of the experiment (12 h to 48 h) indicated that layer
embryos had significantly lower mean heart rate than broiler embryos after 12 h (f HB =
176+7 bpm, fHL = 147+7 bpm; Tukey‟s, q = 4.27, p = 0.004) and 24 h (f HB = 185+5 bpm,
fHL = 167+3 bpm; Tukey‟s, q = 4.22, p = 0.004; Fig. 4.4). However this difference
between broiler and layer fH was no longer significant at 48 h (f HB = 205 + 10 bpm, fHL =
187 + 5 bpm). As expected, the mean heart rate of cultured embryos linearly increased
significantly over a period of 48 hours after the start of the experiment (F (2, 154) = 13.42,
p < 0.001).
Twelve hours from the start of the experiment, yolk environment did not
significantly impact fH of broiler or layer embryos (Fig. 4.4). However, after 24 h the fH of
broiler embryos cultured on layer yolk was significantly lower than that of broiler
embryos cultured on broiler yolk (Tukey‟s, q = 3.44, p = 0.02). Additionally at this time,
fH of broiler and layer embryos exposed to broiler yolk were significantly different from
one another (BB > LB; Tukey‟s, q = 3.04, p = 0.03). However, by 48h post transfer,
mean heart rates of BBs and LBs were no longer statistically different, while mean heart
100
rates of broiler embryos continued to be significantly affected by yolk type (BB > BLs;
Tukey‟s, q = 3.16, p = 0.04). Collectively, these results indicate that broiler embryonic f H
was more sensitive to a breed-specific change in yolk environment than was layer f H,
but only after 24 h of exposure to the foreign yolk environment.
Yolk Hormones
The average yolk T3 concentration in broiler eggs (0.46 pg/mg + 0.22) was
significantly lower than the T 3 concentration detected in layer eggs (1.05 pg/mg + 0.18;
Tukey‟s, q = 3.61, p = 0.02; Fig. 4.5). In contrast, the average yolk TE concentration in
broiler eggs (4.63 pg/mg + 2.02) was significantly higher than the concentrations
detected in layer eggs (3.32 pg/mg + 1.92; Tukey‟s, q = 2.99, p = 0.04). Pearson
product moment correlation analyses revealed that yolk TE and T3 concentrations did
not significantly covary among layer and broiler eggs (r = -0.39, p = 0.52).
Discussion
Body Mass
In ovo, embryonic body mass of broiler embryos and layer embryos differs
significantly, with broiler embryos attaining significantly higher wet and dry body masses
than layer embryos from embryonic days 12 to 20 (Pal et al., 2002; Sato et al., 2006).
The present study extended these observations into earlier development by showing
that after only 64 and 88 h of development on native yolks, broiler embryos were ~3 mg
(wet mass) heavier than layer embryos (Fig. 4.1A, B). Interestingly, the exchange of
embryos and yolks between the two breeds induced significant perturbations in the
natural growth rate of layer embryos, but less dramatic changes in broiler embryo
growth over this early period of development.
101
Sato et al. (2006), suggests that
differential growth rates between the two breeds is the result of differential rates of yolk
lipid absorption and metabolism between the two breeds in late embryonic
development. Genotypic differences between the two breeds have been implicated in
the differential physiology that contributes to growth rate differences among chicken
breeds (Li et al., 2005, 2006). The present study suggested that, prior to embryonic day
4, when yolk mass is not yet a limiting factor in growth, genotypic differences between
the broiler and layer embryos are not the sole determining factor in the breed-specific
mass differences. The question remains as to whether the observed effect of yolk
environment on embryonic body mass is due to differences in lipid concentrations
between the two yolk types, or whether it is due to yolk factors that act to modulate
gene expression which in turn impacts embryonic physiology leading to increased
growth rate.
Development Rate
The comparison of broiler and layer embryo developmental stage as a function of
developmental time has rarely been investigated. Boerjan (2005) suggests that at ~50
h of embryonic development, broiler embryos were more mature (~HH14) than layer
embryos (~HH13), while observations by Yoneta et al., (2007) that broiler embryos
hatched approximately 5 hours later than layer embryos suggests that, late in
embryonic development, broiler embryos may be developmentally delayed with respect
to layer embryos.
The results of the present study show that layer embryos were
significantly more mature (~HH12) than broiler embryos (~HH11) only after 40 h of
development, in ovo (Fig. 4.2).
The paucity of studies dealing with differences in
102
development rate among early-stage embryos begs for further investigation of this
aspect of early development before interpretations can be made.
After 12 h of development on native or foreign yolk, the initial difference in
embryo HH stage between layers and broilers no longer existed (Fig. 4.2). After 24 h of
development, a difference in HH stage between layer embryos exposed to layer yolk
and layer embryos exposed to broiler yolk was detected.
Collectively, these data
strongly suggest that yolk environment plays a significant role in determining the
development rate of embryos. Moreover, the nature of the effects induced by yolk
environment was breed-specific; broiler yolk acted to accelerate development rate of
layer embryos, whereas layer yolk had no apparent effect on the development rate of
broiler embryos. This phenomenon paralleled that observed for body mass (Fig. 4.1),
thus indicating that the increase in body mass of layer embryos cultured on broiler yolk
was likely a function of accelerated development rate rather than increased tissue mass
per se.
The difference in development rate detected at 24 h between layers cultured on
layer or broiler yolk was short-lived, with this effect disappearing by 48 h of incubation
on culture dishes. These occurrences may be explained by a true “critical window” at
40 h to 64 h in chicken development where yolk environment is crucial in determining
embryonic development rate, but has no long-lasting role in shifting the trajectory of this
trait during early development. An alternative explanation is that stage determination
based on Hamburger & Hamilton (1951) does not allow for enough resolution to capture
subtle shifts in embryonic development rate that may be present after 48 h of incubation
on culture yolks.
103
Mass-specific Oxygen Consumption
This is the first comparison of mass-specific oxygen consumption between broiler
and layer embryos during the first 3-4 days of embryonic development. Mass-specific
recorded for 3-4 day embryos (HH17-HH21) in the present study were comparable
to those recorded for layer embryos described by Burggren et al. (2000). Late-stage
broiler embryos (embryonic days 12 to 18) display lower mass-specific
than do layer
embryos of the same age (Sato et al., 2006), but results of the present study suggest
that layer and broiler embryos cultured on native yolks did not differ in embryonic massspecific
. Moreover, yolk environment did not have a great impact on the mass-
specific
of 64-96 h broiler or layer embryos (Fig. 4.3). It is possible that this period
of development was not sufficient to capture potential regulation of mass-specific
metabolic rate via yolk factors. Unfortunately, the culture methods used in this study did
not allow for the assessment of mass-specific
beyond ~88 h of age.
Heart Rate
Besides genotype, many environmental factors significantly influence heart rate
during development. Abiotic factors such as temperature, oxygen availability and light
intensity affect heart rate in chicken embryos, although the nature of the effect is agedependent (Gimeno et al., 1966; Tazawa and Nakagawa, 1985; Chan and Burggren,
2005). In late-stage avian embryos, heart rate is a function of oxygen partial pressure
within the air-cell, which is largely determined by egg shell conductance under normoxic
ambient conditions (Tazawa et al., 2005; Christensen et al., 2006). However, Yoneta et
al. (2005) reports that broiler embryos display significantly faster heart rates than do
layer embryos at embryonic days 10 to 14, and that these differences were not due to
104
differences in shell conductance between the two breeds.
This suggests that egg
factors other than the shell may influence early cardiac development. Here I show that
in the absence of maternally-derived egg shell and albumen, yolk plays a direct role in
ontogeny of heart rate in early-stage chicken embryos.
It appears that 12 h of
development on foreign yolk did not produce significant changes in the embryos‟ f H, but
that 24 h and 48 h of exposure to layer yolk caused broiler embryos‟ f H to be
significantly slower than fH of broilers cultured on native yolk (Fig. 4.4). Interestingly, the
heart rate of layer embryos developing on broiler yolk was higher than that of layer
embryos cultured on native yolk, while the heart rate of broiler embryos on layer yolk
were similar to that of layer embryos cultured on native yolk. This suggests that the
composition of broiler yolk accelerates heart rate, while layer yolk composition
decelerates heart rate, and that this difference in yolk composition may play a role in the
previously reported late-stage heart rate differences between the breeds (Yoneta et al,
2006).
The chronotropic change in broiler embryos due to a change in yolk environment
was not accompanied by any significant changes in body mass or development rate.
As discussed earlier, body mass and development rate of layer embryos were
influenced by the breed-specific changes in yolk environment. Here, I show that the
heart rate of broiler embryos, but not that of layer embryos, was significantly altered by
breed-specific changes in yolk environment. The non-concordance of the effects of yolk
type among these traits is interesting, but not enigmatic. Early in development, chicken
embryo growth (characterized by eye diameter and vasculature growth) and metabolic
rate is not related to cardiac output or heart rate (Burggren et al, 2000; Burggren et al,
105
2004). Thus, in the present study, it can be assumed that yolk-induced perturbations in
heart rate, mass-specific oxygen consumption and growth rate are independent of one
another. However, whether changes in heart rate this early in development contributes
to heart rate regulation, metabolic rate, development rate or growth rate later in
development needs further investigation.
Yolk Hormones
Yolk environment is defined by a multitude of macromolecules which have been
shown to be physiologically relevant to the developing offspring (hormones, lipids,
carbohydrates, carotenoids; Schwabl, 1993; McNabb and Wilson, 1997; Royle et al.,
2001; Saino et al., 2003; Sergiusz et al., 2005). The current results point to these yolk
factors as likely contributors to the determination of morphological and physiological
phenotype of the early developing embryo. Yolk hormones, which have been shown to
be highly variable among and within avian species (Schwabl, 1993; Hahn et al., 2005))
are likely candidates for these observed effects. The present study has characterized
the profiles of two yolk hormones, T 3 and TE, in broiler and layer eggs to elucidate the
potential underlying mechanisms of yolk-induced changes in early embryo morphology
and physiology. In Chapter 2, T 3 increased embryonic heart rate in a dose-dependent
manner, while TE had relatively little effect on heart rate, and neither hormone had an
effect on development rate over the course of the first 1- 4 days of embryonic
development. Since layer yolk induced bradycardia in heart rate of broiler embryos, and
broiler yolk induced slight increases in layer embryos, it would be expected that yolk T 3
levels would be lower in layer yolks than broiler yolks. However, yolk hormone analyses
indicated that broiler yolk T 3 concentration was significantly lower than the yolk T 3
106
concentration of layer eggs by ~2-fold (Fig. 4.6). In light of this finding, it was unlikely
that yolk T3 is directly responsible for the observed chronotropic effects of chicken yolk.
In contrast to yolk T3 concentration, yolk TE concentration was significantly greater in
broiler yolks than layer yolks. Because there was no effect of TE on heart rates of early
layer chicken embryos (Chapter 2), it is unlikely that the significant difference in TE
between the two breeds was responsible for the chronotropic responses observed in
layer embryos cultured on broiler yolk, or vice versa.
These conclusions were made under the assumption that, with respect to heart
rate and development rate, broiler embryos and layer embryos respond to T 3 and TE in
a similar fashion. It was likely the case that the yolk effects seen in the current study
was a result of the complex interaction of breed-specific yolk factors and breed-specific
genotype. Also, T3 and TE are only two of numerous yolk hormones that have been
implicated in modulating development (Groothuis et al., 2005).
Thus, a complete
analysis of yolk hormones and other yolk factors are needed to better understand the
mechanisms underlying these findings.
Egg Components Mass and Egg Mass
Egg components mass and egg mass of broiler and layer chicken eggs in this
study were comparable to those reported previously (Finkler et al., 1998; Sato et al.,
2006; Yoneta et al, 2007). Similar to Sato et al. (2006), the current results show that
broiler eggs, although not different in mean mass to layer eggs, possessed significantly
larger yolk mass than layer eggs. This difference in yolk mass between the two breeds
was compensated for by significantly smaller albumen mass in broiler eggs when
compared to layer eggs (Fig 4.6).
107
Yolk mass and egg mass significantly predicts hatchling mass, hatchling maturity
and offspring performance in oviparious species (Sotherland and Rahn, 1987; Sinervo
and Huey, 1990; Dzialowski and Sotherland, 2004). In the current study, it appears that
the larger yolk mass of broiler eggs correlated with larger body mass of those embryos
cultured on broiler yolk, while the relatively smaller yolk mass of layer eggs correlated
with the relatively smaller body mass of those embryos cultured on layer yolk (Fig. 4.1).
This suggested that the yolk mass may be useful in predicting embryo mass during
early development (HH 9-14), a period when yolk volume per se does not play a role in
limiting body mass.
Summary
Inter-breed differences between broiler and layer chickens have traditionally
been attributed largely to direct genetic effects.
However, during early embryonic
development (first four days after incubation), chicken embryos rely heavily on egg
components for growth and development, which strongly implicates maternal effects in
the development of breed-specific phenotypes.
Unfortunately, few studies have
investigated the role of maternally-derived yolk environment in determining broiler and
layer phenotypes during this early period. The current study sheds light on the role of
the yolk environment in determining the well-characterized phenotypes of the layer
chicken and the broiler chicken, each of which have been artificially selected for traits
serving different purposes, and show genotypic differences.
Moreover, this study
revealed a complex interaction of breed-specific genotype and breed-specific yolk
environment (possibly yolk hormones) early chicken development. These novel findings
have crucial implications in the significance of maternal effects and/or epigenetic
108
processes on early embryonic heart rate, growth rate, oxygen consumption and
development rate of vertebrates.
In Chapter 5, I now consider non-avian parental effects by examining zebrafish
populations exposed to variable levels of oxygen availability.
109
A
Wet Body Mass (mg)
25
LB
BB
LL
20
BL
LB > LL (q=3.08, p=0.03)
15
10
LB > LL (q=3.00, p=0.04)
0
0
Dry Body Mass (mg)
B
10
20
30
40
50
60
2.0
1.8
LB
1.6
BB
1.4
LB > LL (q=3.39, p=0.02)
LL
BL
1.2
0.0
0
10
20
30
40
50
60
Time Elapsed (h)
FIGURE 4.1 Wet (A) and dry (B) body mass of broiler and layer embryos cultured on
broiler or layer egg yolk for a period of 24 and 48 hours after the start of experiments.
Group designations (BB, BL, LB and LL) refer to the embryo type and yolk type,
respectively, for each group. A box indicates no significant difference among the
groups. N’s > 7 for each group at each time points. Data are presented as mean +
S.E.M.
110
Developmental Stage (HH)
20
18
16
LB > LL (q=3.40, P=0.02)
14
12
LB > BB (q=2.97, P=0.04)
0
-10
0
10
20
30
40
50
Time Elapsed (h)
FIGURE 4.2 Developmental stage of broiler and layer embryos cultured on broiler or
layer egg yolk for a period of 12, 24 and 48 h after the start of experiments. Group
designations (BB, BL, LB and LL) refer to the embryo type and yolk type, respectively,
for each group. Significant differences between groups are noted at each time point.
N’s > 7 for each group at each time points. Data are presented as mean + S.E.M.
111
A
Mass-specific VO2 (ml g-1 h-1)
25
.
.
VO2 = -0.75M + 18.79
20
all
(r2 =0.61, N=42, P<0.001)
15
10
5
0
0
B
5
Mass-specific VO2 (ml g-1 h-1)
25
.
10
.
VO2
BL
15
20
25
= -1.01M + 122.10
2
(r =0.68, N=10, P=0.003)
20
.
= -0.62M + 17.39
VO2
LB
2
(r =0.43, N=13, P=0.02)
15
.
VO2
10
2
BB
= -0.62M + 16.85
(r =0.79, N=8, P=0.003)
.
5
VO2
LL
= -0.72M + 17.50
2
(r =0.61, N=11, P=0.004)
0
0
5
10
15
20
25
Body Mass (mg)
FIGURE 4.3 Mass-specific oxygen consumption of all cultured embryos (A), and broiler
and layer embryos cultured on broiler or layer egg yolk (B) as a function of body mass.
Group designations (BB (circle), BL (triangle), LB (square) and LL (diamond)) refer to
the embryo type and yolk type, respectively, for each group.
112
220
fH (bpm)
200
180
BB
BL
160
LB
LL
140
0
0
10
20
30
40
50
Time (h)
FIGURE 4.4 Mean heart rate of broiler and layer embryos cultured on broiler or layer
egg yolk for a period of 12, 24 and 48 hours. Group designations (BB, BL, LB and LL)
refer to the embryo type and yolk type, respectively, for each group. A box indicates no
significant difference among groups. N’s > 6 for all groups at each time point, except
BB at time 48, n = 3; and BL at time 48, n = 2. Data are presented as mean + S.E.M.
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6
Concentration (pg mg-1)
Broiler
Layer
(12)
*
P < 0.05
5
(12)
4
3
2
1
*
P < 0.05
(4)
(3)
0
T3
TE
Yolk Hormone
FIGURE 4.5 Yolk T3 and TE concentrations of broiler and layer eggs. An asterisk
indicates significant difference between layer and broiler yolk hormone concentration.
Numbers above bars indicate n for each group. Data are presented as mean + S.E.M.
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Egg Components Mass (g)
50
40
30
20
10
Albumen
Yolk
Shell
0
45
50
55
60
65
70
Egg Mass (g)
FIGURE 4.6 Egg components mass of broiler and layer eggs as a function of egg
mass. Filled symbols represent broiler egg components and unfilled circles represent
layer egg components. Lines through data points indicate best fit regression of each
egg components‟ mass and egg mass. Regression models for layer egg components
and egg mass are as follows: Myl = 0.24Mel + 2.43; r2 = 0.33; Mal = 0.74Mel – 6.97; r2 =
0.82; Msl = 0.08Mel + 1.79; r2 = 0.31. Regression models for broiler egg components
and egg mass are as follows: Myb = 0.28Meb + 2.21; r2 = 0.48; Mab = 0.78Meb – 11.13; r2
= 0.084; Msb = 0.07Meb + 1.85; r2 =0.41.
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CHAPTER 5
INFLUENCE OF PARENTAL HYPOXIA EXPOSURE ON STRESS SURVIVAL
PHYSIOLOGY OF ZEBRAFISH LARVAE
Introduction
In response to perturbations in the environment, adults of a wide range of
vertebrate species show beneficial changes in parental investment and care that
promote advantageous offspring phenotype (Munkittrick and Dixon, 1988; Meyer and Di
Giulio, 2003; Mondor et al., 2005). This is traditionally known as an “advantageous
environmentally-induced parental effect” or an “indirect genetic effect” (Lacey, 1998).
Examples include cotton aphids that produce more winged offspring when exposed to
predator attacks, presumably to enhance the ability of their offspring to escape
predation (Mondor et al., 2005).
Offspring of killifish and feral white sucker fish of
polluted environments produce offspring that are more resistant to the specific pollutant
(Munkittrick and Dixon, 1988; Meyer and Di Giulio, 2003). These lines of evidence
exemplify a strong link between the stressors experienced by the parents and the
increased fitness of the offspring to survive these same stressors.
Currently, one of the most prevalent stressors in aquatic ecosystems is the
depletion of oxygen due to human anthropogenic activities (Diaz, 2001). Thus there is
an urgent need to characterize the effects of parental hypoxia exposure on offspring
hypoxia survival in aquatic animals. Hypoxia exposure in adult fish largely results in
reduced fitness due to low fecundity, decreased mating rates, and low survivorship of
offspring (Wu et al., 2003). Adult carp, for example, exposed to moderate levels of
hypoxia suffered disruption of the endocrine systems that resulted in lowered
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hatchability of the offspring as well as lethal mutation phenotypes in offspring (Wu et al.,
2003).
Beyond the gross morphological effects of parental hypoxia exposure on
offspring phenotype, little is known of the effects on the offspring‟s physiology; more
specifically, of the effects on the offspring‟s ability to cope with hypoxic stress through
physiological responses as part of overall acclimatization. The only line of evidence that
points to a possible advantageous parental effect in the face of parental hypoxia
exposure is that, in D. melanogaster, offspring of hypoxia-reared parents, raised in
normoxia developed larger tracheal systems (Henry and Harrison, 2001).
Larger
tracheal system is suggestive of increased hypoxia resistance due to increased
respiratory surface area. However the potential for greater resistance to hypoxia in
those with larger tracheal systems was not directly tested.
I hypothesized that aquatic animals, such as fish, express advantageous
hypoxia-induced parental effects, and that these effects translate into increased
resistance to hypoxia in the larvae of the subsequent generation.
Mature and
developing zebrafish have been reported to be sensitive to the effects of chronic and
acute hypoxia, and these responses have begun to be characterized on a behavioral,
physiological, biochemical and molecular level (Pelster and Burggren, 1996, Padilla and
Roth, 2001; Rees et al., 2001; Jacob et al., 2002; Ton et al. 2003). The on-going
investigation of the impact of variable oxygen levels in zebrafish makes it a valuable
model for the investigation of trans-generational effects of hypoxia.
In the present
study, I assessed not only the hypoxic survival response but also the thermal tolerance
of the larval progeny of adult zebrafish exposed to hypoxic conditions for controlled
periods of time. Thermal stress elicits similar molecular and biochemical changes as
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hypoxic stress (Iwama, 1999), so the dual study of hypoxic and thermal stressors would
allow for the assessment of the potential mechanisms of hypoxia-induced parental
effects. I also assessed the body length of the progeny, and the clutch size and egg
components of the hypoxia-exposed zebrafish to better understand how hypoxiainduced changes of maternally-derived factors may impact offspring phenotype.
Materials and Methods
Care and Maintenance
Adults. Wild-type zebrafish (Danio rerio) obtained from Ekkwill Farms, FL, were
housed in glass aquariums at a density of 10 fish/13 liters. Adults were exposed to 2
weeks of laboratory conditions prior to the start of experiments. They were fed brine
shrimp and dry flake food twice a day during non-breeding periods, and 3 times per day
5 days prior to breeding. Care was taken to feed equal amounts of food to fish held in
separate tanks. Temperature inside the tanks was maintained at 27 + 0.5°C by a
submersible water heater. Light cycle was set at LD 14 h:10 h.
Embryos and larvae.
Embryos (~70 per 135 mm x 10 mm Petri-dish) were
housed in 95-mm Petri dishes filled with clean, normoxic tank water treated with 0.05%
methylene blue (14:10 h light:dark cycle). After 5 days post-fertilization (dpf), larvae
were transferred to 1 L containers at a density of ~150 larvae/L. Air saturation of the
water in each container was maintained by bubbling ambient air into the water.
Beginning at 6 days post fertilization (dpf), larvae were fed dry powder food (TetraMin,
Baby Fish Food “E” for egglayers) 3 times daily. Brine shrimp were added to feedings
beginning at 16 dpf. All embryos and larvae were housed in a temperature-controlled
incubation room maintained at 27 + 0.5°C .
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Water PO2. Normoxia was maintained by constantly running room air through a
sponge filter located at the bottom of the tank. Hypoxia was maintained in a tank by
constantly running a gas mixture of 13% oxygen and 87% nitrogen through a sponge
filter located at the bottom of the tank. Tanks were fitted with a gas-tight lid equipped
with a circular opening small enough to create positive air pressure inside the tank that
only allowed gas from the air-stone to escape.
Routine breeding.
During the holding period of at least 2 weeks, fish were
regularly bred to increase the likelihood of successful breeding during experiments.
Three males and three females were placed in breeding containers filled with clean,
normoxic tank water the evening prior to the morning of egg collection.
Breeding
containers consisted of a 1-liter rectangular container that held a smaller rectangular
container with a mesh bottom. The bottom of the inner container rested about 0.5
inches above the bottom of the outer container to allow for eggs to fall to the bottom,
and prevent them from being eaten by the adult fish. Breeding tanks were maintained
in a 27°C incubator room during the breeding period. Four hours after the lights came
on the next morning, fish were allowed to breed, and were then returned to aquatic
holding tanks.
Experimental Design
Breeding pairs that produced > 100 eggs during routine breeding were assigned
to one of two experimental conditions, hypoxia-exposure or normoxia-exposure. Prior
to placement into experimental tanks containing hypoxic or normoxic water, pairs were
bred, and the eggs discarded, to ensure that the majority of pre-existing eggs were
eliminated before experiments began. Twelve breeding pairs were assigned to each
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experimental group. The evening prior to the start of an experiment, breeding pairs
were placed in individual breeding tanks containing normoxic water overnight. Breeding
in normoxia ensured that eggs of both hypoxia and normoxia groups were exposed to
the similar conditions. A tank divider was used to keep the female and male separated
until the start of the light cycle to control for spontaneous breeding during the dark cycle.
At the start of the light cycle, the divider was removed, and pairs were allowed to breed
for ~3 h before eggs were collected and rinsed in clean tank water.
Both morphological and physiological aspects were assessed in zebrafish larvae
whose parents (P) were exposed to 1, 2, 3 or 4 wks of hypoxia (HP-1, HP-2, HP-3 or
HP-4, respectively) or normoxia (NP-1, NP-2, NP-3 or NP-4). Hypoxic survival time and
body length (BL) of these larvae were assessed at 6, 9, 12, 15 and 18 dpf.
For
measures of hypoxia survival time and BL, larvae were kept in their respective clutches
for the assessment of inter-clutch variability within HP and NP groups. Each clutch, at
every period of parental exposure to experimental conditions (1, 2, 3 or 4 wks),
represented an entirely different breeding pair.
In a separate experiment designed to assess the effects of hypoxia exposure on
the fecundity and egg characteristics of fish exposed to hypoxia or normoxia, cohorts of
5 females and 5 males were exposed to experimental conditions for 1, 2, 6 and 12
weeks to assess fecundity (eggs per clutch), and whole egg, yolk and perivitelline
volume of collected eggs. At each point during the course of exposure to experimental
conditions, breeding pairs were randomly selected from each experimental group and
returned to assigned tanks for subsequent exposure. Thus, groups over time represent
the same cohort of fishes, although egg clutches were not obtained from the same
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breeding pairs, or from every pair at each exposure time. Also, during this experiment,
after 12 wks of treatment exposure, the critical thermal maxima and minima of NP and
HP larvae were recorded at 6, 9, 12, 15, 18, 21 and 60 dpf to assess the thermal
tolerance of these two groups.
Larval Body Length
Standard body length was defined as the distance from the tip of the snout to the
rear center of the tail, and was assessed in larval zebrafish using an imaging system
consisting of a compound microscope with attached digital camera and ImagePro Plus
data analysis software (v. 4.1, Media Cybernetics).
Fecundity and egg, yolk and perivitelline volume
Immediately after a 3 h breeding period, eggs were collected from breeding tanks
and counted. Images of the eggs were captured using a high-speed camera (Model
JE3010, Javelin Electronics). Egg, yolk, and perivitelline volume were assessed by
making measurements of egg and yolk diameter from light microscopy images of 20X
magnification (Image Pro Plus v. 4.1). Egg and yolk volumes were derived using the
equation,
, where r is the radius of the component of interest. Perivitelline
volume was calculated by subtracting yolk volume from egg volume.
Hypoxia Survival
Time to Loss of Equilibrium to Hypoxia (T LOE(H)). The time that it takes for fish to
lose equilibrium when exposed to certain stressors is an indication of its resistance or
tolerance to that stress. Loss of equilibrium has been considered a reliable and valid
measure of “ecological death” in many fish species, and has been used in studies as an
alternative to physiological death (Beitinger et al., 2000).
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Zebrafish larvae (6, 9, 12, 15, 18, 21 dpf) were placed individually in an airtight
acrylic box (10cm x 10cm x 5cm) filled with extremely hypoxic fish water (~30mmHg
O2). Timing started immediately upon hypoxic exposure, and ended upon the larva‟s
inability to recover from loss of equilibrium (failure to right itself) for at least a 3 sec
period. Three sec were subtracted from the total time to account for the time taken to
determine loss of equilibrium. Embryos younger than 6 dpf were not free swimming,
and do not display clearly observable loss of equilibrium, and thus were excluded from
experiments.
Critical Thermal Maxima and Minima (CTMax and CTMin).
The critical thermal method (CTMax and CTMin) is frequently used to estimate
the scope of thermal tolerance (aka thermal scope) of an organism (Herrera et al., 1998;
Ford and Beitinger, 2004). The method directly quantifies the temperatures at which an
aquatic organism loses equilibrium. Zebrafish larvae (6, 9, 12, 15, 18 and 21 dpf) and
juveniles (60 dpf) were placed in 100-ml glass beakers filled with fresh normoxic tank
water. The beakers were then placed in a temperature-controlled water bath (Fisher
Isotemp 1028P). Fish were kept in the beakers < 5 min before the temperature of the
water was increased (CTMax) or decreased (CTMin) at a rate of 0.3 + 0.1°C min-1. This
rate of temperature change is recommended for small fish since it allows for an
insignificant time lag between water temperature and body temperature (Beitinger et al.,
2000).
The temperature at which the larva was unable to recover from loss of
equilibrium for at least 3 sec was recorded as the CTMax or the CTMin. Since the rate
of water temperature change was variable (+1°C min-1) from trial to trial, each trial
consisted of 3 larvae from each experimental group. Prior to 12 dpf, CTMin of larvae
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were not obtainable due to the difficulty of assessing loss of equilibrium at extreme cold
temperatures. Therefore, CTMin was assessed in larvae at 12, 15, 18, 21 and 60 dpf.
The thermal tolerance of larval zebrafish (6 to 21 dpf) is unknown, thus the
thermal tolerance of NP-12 and HP-12 juvenile (60 dpi) zebrafish were assessed to
determine the validity of the protocol by comparison to adult zebrafish studies reporting
thermal scope.
Statistical Analyses
Larval hypoxic survival time (TLOE(H)) and body length were subjected to factorial
three-way analyses of variance (ANOVA) with respect to treatment group (NP or HP),
parental exposure period (1, 2, 3, and 4 wks), and developmental age (6, 9, 12, 15 and
18 dpf). Subsequent separate two-way analyses with respect to treatment group and
development measured as dpf were performed for each parental exposure period.
Also, inter-clutch analyses of variables were carried out using two-way ANOVA with
respect to clutch and developmental time for each exposure period. In cases where an
analysis of interactions could not be performed due to unequal groups within factors,
one-way ANOVAs were performed for each developmental time period. Subsequent
Tukey‟s post-hoc analyses were used to detect differences between treatment groups
with respect to treatment exposure times.
Variables of fecundity, whole egg volume and egg component volumes were
analyzed in a similar fashion using two-way ANOVAs on factors of treatment group
(normoxia or hypoxia-exposed) and treatment exposure times (1, 2, 6, or 12 wks). A
two-way ANOVA with respect to treatment group (NP and HP) and developmental time
(6, 9, 12, 15, 18, 21 and 60 dpf) were performed on variables of CTMax and CTMin.
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Subsequent Tukey‟s post-hoc analyses were used to detect differences between
treatment groups. For all analyses, if normality and/or equality of variances were not
achieved, data were ranked prior to statistical analyses. Statistical significance was
assigned a value of p < 0.05. Data were expressed as means + S.E.M.
Results
Fecundity and Egg Components Volume
The fecundity of adult zebrafish exposed to 1, 2, 6 or 12 weeks of hypoxia or
normoxia is depicted in Figure 5.6.
After 1 week of exposure to experimental
conditions, mean fecundity of hypoxia-exposed fish was not significantly different from
that of normoxia-exposed fish, but that after 2, 6 and 12 wks of exposure, hypoxiaexposed fish laid fewer eggs per clutch than normoxia-exposed fish (q = 2.96, p =
0.048; q = 3.06, p = 0.042; q =2.99, p = 0.046, respectively).
The egg, yolk, and perivitelline volumes of the eggs produced by the
aforementioned zebrafish are depicted in Figure 5.7. Normoxia-exposed fish produced
eggs with significantly larger egg volume and perivitelline volume than hypoxia-exposed
fish after 1 and 2 wks of exposure (q‟s > 4.62, p‟s < 0.001). However, this difference
disappeared after 6 an 12 wks of exposure.
Mean yolk volume was not different
between the two treatment groups at any exposure period.
Larval Body Length
Mean larval body length significantly increased as a function developmental time
for both NP and HP larvae (p‟s < 0.05 ; Fig. 5.4A-D). However, this increase in mean
body length differed in magnitude between NP and HP larvae, with HP larvae displaying
significantly longer mean body length than NP larvae, at variable points during larval
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development, over the course of the four parental exposure periods (1, 2, 3 and 4 wks).
After 1 wk of exposure to experimental conditions, HP larvae were longer than NP
larvae at 6, 15, and 18 dpf (q = 2.147, p = 0.03; q = 3.62, p < 0.001; t = 4.70, p < 0.001,
respectively; Fig. 5.4A). As parental exposure period increased to 2 wks, the difference
in mean body length between NP and HP larvae was lost at 6dpf, but remained
significant at 15 (q = 3.90, p = 0.006) and 18 dpf (q = 6.58, p < 0.001; Fig. 5.4B).
Exposure periods of 3 and 4 wks caused further dampening of mean body length
difference between NP and HP larvae, with HP larvae significantly longer than NP
larvae only at 18 dpf (3 wk exposure (Fig. 5.4C); q = 3.24, p = 0.002), and 15 dpf (4wk
exposure (Fig. 5.4D); q = 2.57, p =0.011).
Significant inter-clutch variation within NP and HP larvae groups were detected
for all exposure periods (P‟s < 0.05; Fig. 5.5A-D). Among NP-1 clutches, there were no
significant differences in mean body length at any time in development, while among
HP-1 clutches, mean body length of one clutch was significantly different than the rest
of the clutches at 9, 12, 15 and 18 dpf (q‟s > 5.06, p‟s < 0.01; Fig. 5.5A). Similarly, the
mean body length of HP-2 clutches were significantly different at 6, 9, 15, and 18 dpf
(q‟s > 3.85, p‟s < 0.033), while NP-2 clutches were not different from one another (Fig.
5.5B). This effect remained significant at 9 dpf for HP-3 clutches (q = 4.21, p = 0.02),
and at 6, 12, and 15 dpf among HP-4 clutches (q‟s > 4.94, p‟s < 0.01). However, there
were no differences detected in mean body length among NP-3 or NP-4 clutches at any
developmental time (Fig. 5.5C and D).
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Hypoxia Survival
The sensitivity of zebrafish larvae to extreme hypoxia (30 mmHg O 2) increased
over the course of early development, with larvae showing a ~2-fold decrease in time to
loss of equilibrium from 6 to 18 dpf (p < 0.001; Fig. 5.1). Subsequent two-way ANOVAs
indicated that there was a significant effect of time and treatment for at each parental
exposure period (p‟s < 0.05). In particular, after 1 wk of parental exposure to hypoxic or
normoxic conditions, 9 and 12 day old HP larvae displayed shorter mean T LOE(H) than
NP larvae (Tukey‟s, q = 2.93, p = 0.038; q = 2.85, p = 0.04, respectively; Fig 5.1A).
The increase in hypoxic sensitivity during development was reversed, however,
when parental fish were reared in treatment conditions for 2, 3 and 4 wks (Fig. 5.1B, C
and D). After 2 wks of treatment, hypoxia-exposed zebrafish produced offspring that
were more capable of coping with hypoxia stress than the offspring of normoxiaexposed fish, at 9 (q = 3.90, p = 0.006) and 18 dpf (q = 3.25, p = 0.022). After 3 and 4
wks of exposure, 12 and 18 day old HP larvae were better able to combat hypoxia
stress than NP larvae (q = 5.47, p < 0.001; q = 3.03, p = 0.034).
A comparison of mean TLOE(H) among clutches revealed high variability both
among and between clutches of NP and HP larvae (Fig. 5.2). Although there was a
significant effect of treatment (parental normoxia or hypoxia exposure) on larval mean
TLOE(H) after 1 wk of exposure (Fig. 5.1A), inter-clutch analyses revealed no significant
differences in mean TLOE(H) among clutches (Fig. 5.2A). In contrast, after 2, 3 and 4 wks
of exposure, HP clutches showed significant variation in mean T LOE(H), with one or two
clutches appearing to drive the significant difference in mean TLOE(H) observed between
NP and HP groups (compare Fig. 5.1B, C and D and Fig.5.2B, C and D). After 2 wks of
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exposure, there was a significant difference in hypoxia sensitivity between the two HP
clutches at 8dpi, while no inter-clutch differences were observed for NP larvae (q = 4.80,
p = 0.004; Fig. 5.2B). Similarly, after 3 wks of parental treatment exposure, significant
differences in TLOE(H) only existed between HP clutches at 9 dpf (q = 4.37, p = 0.01).
After 4 wks (Fig. 5.2D), significant differences in TLOE(H) among HP clutches existed at
12 dpf (q = 4.92, p = 0.01), while a significant difference was detected within NP
clutches at 9 dpf (q = 4.96, p = 0.008).
CTMin and CTMax
In an experiment separate from that of larval hypoxia survival, thermal tolerance
of larval (6 to 21 dpi) and juvenile (60 dpi) offspring of hypoxic- or normoxic-exposed (12
wks) adult zebrafish were characterized by the assessment loss of equilibrium in the
face of increasing (CTMax) or decreasing (CTMin) temperatures.
There were no
observable difference in thermal tolerance between the HP-12 larvae and the NP-12
larvae at any time in development (p‟s < 0.05; Fig. 5.3). In general, over the course of
development from 6 to 60 dpf, zebrafish showed an increase of thermal tolerance.
Discussion
Larval Hypoxia Survival and Normal Development
Hypoxia resistance is defined by an animal‟s ability to combat the damaging or
lethal effects of low oxygen levels by maintaining normoxic oxygen levels in tissues
despite low ambient levels. Hypoxia tolerance, in contrast, is defined by an animal‟s
ability to cope with low oxygen partial pressure in tissues, which can include simple
tolerance of hypoxia. In euryoxic fish, many factors have been shown to contribute to
hypoxia resistance such as oxygen affinity of hemoglobin, ventilation rate, cardiac
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output, diffusion coefficient of respiratory surfaces, and/or the differential shunting of
oxygen-rich blood to vital organs (Boutilier and St.-Pierre, 2000).
Mechanisms of
hypoxia tolerance in fish, in contrast, are largely attributable to the switch to glycogen
stores for anaerobic respiration, down-regulation of metabolic rate, and/or the induction
of heat shock proteins (Hochachka et al., 1996).
During the early phases of
development when oxygen diffusion is the predominant means of supply to tissues,
zebrafish are likely tolerant of oxygen depletion in the environment, while later in
development, as physiological processes begin to play a more significant role in the
response to lowered oxygen availability, it is likely that resistance and tolerance
responses both determine the animal‟s survival in hypoxia (Pelster and Burggren, 1996;
Padilla and Roth, 2001; Jacob et al., 2002).
As embryos, zebrafish display increased sensitivity to environmental oxygen
depletion as development progresses (Padilla and Roth, 2001; Ton et al, 2003). At the
two-cell stage, zebrafish embryos are able to tolerate anoxic conditions by entering into
a state of suspended animation, with this anoxia tolerance gradually disappearing by
the time of hatching (Padilla and Roth, 2001). Moreover, after 24 h post-fertilization,
zebrafish embryos become increasingly sensitive to environments of 5% oxygen (Ton et
al., 2003). The present study extended these previous findings by demonstrating that
from 6 to 18 dpf, zebrafish larvae sensitivity to severe, acute hypoxia (30mmHg O 2)
gradually increased as development progressed (Fig. 5.1A-D). During this period of
larval development (6 to 18 dpf), zebrafish larvae undergo both qualitative and
quantitative changes in mode of oxygen transport to tissues and oxygen demand of
tissues (Pelster and Burggren, 1996; Jacob et al, 2002; Rombough, 2002). Prior to 12
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days post-fertilization (dpf), oxygen supply to the tissues occurs via diffusion across the
body surface, but as the larva increases in size (~14 dpf), convective circulation
becomes necessary to maintain adequate oxygen supply to tissues (Jacob et al., 2002).
It is during this time that cardioventilatory mechanisms combat the effects of hypoxia
(Jonz and Nurse, 2005; Jonz and Nurse, 2006; Vulesevic and Perry, 2006; Vulesevic et
al, 2006).
It is likely that these developmental milestones in oxygen regulation
contribute to the decline in hypoxia resistance in larval zebrafish observed in the
present study. The decrease in hypoxia resistance and/or tolerance during this period
of transition in the zebrafish larvae is interesting and deserves further study to uncover
the mechanisms (resistance versus tolerance) of hypoxia survival response in larva fish.
Since it remains unknown whether survival during a bout of severe hypoxia in larvae is
primarily due to resistance strategies or tolerance strategies, sensitivity to hypoxia will
be arbitrarily be termed “resistance.”
Larval Hypoxia Resistance and Parental Influences
The present study is the first to characterize hypoxia resistance of the offspring of
zebrafish larvae from adult populations exposed to chronic hypoxia compared to
normoxia. Generally, larvae of hypoxia-exposed adult zebrafish showed significantly
greater tolerance to an acute bout of severe hypoxic stress than did the larvae of
normoxia-exposed zebrafish, as measured by T LOE(H) (Fig. 5.1B, C, D). However, 1
week of parental exposure to hypoxia or normoxia, the offspring of hypoxia-exposed
adults were less resistant to hypoxia than the larvae of normoxia-exposed adults (Fig.
5.1A). This suggests that the underlying mechanisms responsible for the parental effect
of heightened survivability in acute, severe hypoxia in larvae do not manifest until after 1
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week of parental hypoxia exposure. Evidence that gene expression profiles of euryoxic
fish (Gillichthys mirabilis) change significantly depending on the length of time the fish
were exposed to hypoxia (8 h to 6 days) support the current hypothesis (Gracey et al.,
2000). Eight hours after the onset of hypoxia induction, relatively few genes were upregulated while when compared to the number of genes up-regulated at 6 days hypoxia
exposure. However, it is not known whether the genes that are temporally regulated
are involved in the phenomenon observed presently, or if exposure to hypoxia beyond 6
days (as was the case in this study) induces further changes in gene expression.
The effect of parental hypoxia exposure on hypoxia resistance of larval zebrafish
in this study appeared to be largely clutch-dependent, with TLOE(H) being significantly
different among HP clutches, but not NP clutches (Fig. 5.2B-D).
In most cases, the
overall difference between NP and HP groups could be explained by a single highly
deviant HP clutch. Moore et al (2005) show that the effects of hypoxia on zebrafish
cardiovascular parameters were highly variable among families, and suggest that this
variable interaction of genotype and hypoxia is the basis for adaptation and evolution.
The present study demonstrates that, in addition to the highly variable effects of hypoxia
exposure on the cardiovascular system reported by Moore et al (2000), there is a
significant interaction of parental genotype and low oxygen environment that produces
highly variable regulation of progeny stress resistance. Due to the rapid response to
lethal levels of hypoxia (T LOE(H) < 253 s for HP larvae and < 100 s for NP), it is likely that
the mechanism underlying hypoxia resistance differences both between and within HP
and NP groups was associated with perturbations in basal level functioning, basal
protein levels (i.e. constitutive and inducible heat shock protein 70 (Beckmann et a.l,
130
1990)) and/or cardioventilatory responses associated with oxygen regulation, rather
than a difference in the ability to increased or decreased protein expression in direct
response to hypoxia, as seen in long-term, non-lethal hypoxia exposure (Gracey et al.,
2000).
The current findings expand upon earlier reports that hypoxia exposure in the
parental generation induces morphological changes suggestive of increased hypoxia
resistance (i.e. increased tracheal size in D. melanogaster) in the subsequent
generation (Henry and Harrison, 2001). However, further physiological, biochemical
and molecular analyses of the HP and NP larvae are required to understand the
mechanisms that underlie the effects of parental hypoxia on hypoxia resistance in larval
fish.
Thermal Tolerance
The upper thermal tolerance of larval and juvenile zebrafish (60 dpf) in this study
was comparable to those reported for adult zebrafish (Cortemeglia and Beitinger, 2006).
However, the lower thermal tolerance of larval (12 – 21 dpf) and juvenile fish was
approximately twice as high as those previously reported for adults (Cortemeglia and
Beitinger, 2006). This indicates that thermal scope of developing zebrafish is narrower
than that of adults, with the lower limit shifted upwards. However, due to the difficulty of
assessment of loss of equilibrium in response to cold temperatures in larval zebrafish
(they sit at the bottom and do not lose equilibrium), the current estimates of CTMin in
larval and juvenile zebrafish may be not be representative of the actual lower thermal
tolerance.
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Hypoxia exposure induces very similar gene expression profiles to that of
extreme temperature exposure, suggesting an intimate relationship between hypoxic
resistance and thermal tolerance (Airaksinen et al., 1998; Sørensen and Loeschcke,
2001; Sakamoto et al, 2005; Deane and Woo, 2006). Regulatory proteins such as heatshock proteins and hypoxia-inducible factor-1 have been shown to be up-regulated
similarly to hypoxic and thermal stress (Rissanen et al., 2006; Horowitz, 2007; Tetievsky
et al., 2008). Moreover, resistance (or exposure) to one stress has been shown to
confer resistance to a wide range of other stresses. For example, thermal acclimation
in adult C. elegans confers tolerance against not only thermal stress, but also hypoxia
and heavy metal stress (Katschinksi and Glueck, 2003).
In this study, the thermal
tolerances of the offspring (larval and juvenile) of hypoxia-exposed adult zebrafish and
that of offspring of normoxia exposed adult zebrafish were not significantly different (Fig
5.3).
This suggests that the regulation of hypoxia resistance does not necessarily
involve the regulation of genes common between hypoxia and heat shock stress, such
as heat-shock proteins (Iwama, 1999). Recently, Kassahn et al (2007), reported that
although stress responses of coral reef fish brought on by a variety of stressors
(including hypoxic and cold and heat shock) induced changes in common functional
gene groups, hypoxic stress induced widely different changes in expression of individual
genes within these functional groups when compared to cold or heat shock stress. This
lends credence to the hypothesis that the molecular mechanisms that underlie
heightened hypoxia resistance in HP larvae are hypoxia-specific.
132
Larval Body Length
The current findings show that the body lengths of larvae from hypoxic parents
were larger than that of larvae from normoxic parents at variable times in development,
with the effect disappearing and reappearing over the course of the experiment (Fig.
5.4A-D). The majority of differences between the two groups were observed at 15 and
18 dpf, with NP-1 and HP-1 larvae showing an additional difference as early as 6 dpf.
Similar to larval hypoxia resistance, larval body length shows significant inter-clutch
variation within the HP group only. Hypoxia exposure apparently exaggerates the
inherent variation in parental effects pertaining to body size, within a population of fish.
Although the relationship between body size and hypoxia resistance/tolerance is
controversial, body size has clear implications in oxygen regulation (Feder, 1983a, b;
Burleson et al, 2001; Robb and Abrahams, 2003; Ospina and Mora, 2004; Nilsson and
Ostlund-Nilsson, 2008).
Nilsson and Ostlund-Nilsson (2008) suggests that larger
bodied fish are better able to survive severe hypoxia stress due to increased levels of
glycogen available for anaerobic adenosine-triphosphate production, while Robb and
Abrahams (2003) found that large yellow perch (34.07 g) were significantly less tolerant
to extreme hypoxia stress than small individuals of the same species (2.29 g).
However, these reports were based on adult fish, and thus cannot be fully extended to
the current study of developing fish. Because instances of longer body length did not
correlate with instances of increased hypoxia resistance during larval development
(compare Fig. 5.1A-D with Fig. 5.4A-D), it is likely that these two parameters are both
results of parental hypoxia exposure, but are independent of one another. In support of
133
this conclusion, independence of oxygen regulation and body size has been also been
observed in frog larvae (Feder, 1983a, b).
Fecundity and Egg Volume
Egg components and fecundity are largely under maternal control. The ability to
alter these two components of reproduction have been implicated in strategic responses
to environmental conditions to increase fitness of the individual and/or its offspring
(Einum and Fleming 2004; Dziminski and Alford, 2005; Landry et al, 2007). In the
present study, hypoxia-exposed adult zebrafish produced smaller fewer eggs than
normoxic controls only after 2 weeks of hypoxia-exposure (Fig. 5.6 and 5.7). After 1
week of hypoxia-exposure, hypoxia-exposed fish produced smaller eggs but did not
differ in fecundity from normoxia-exposed group, while after 6 and 12 wks of hypoxia
exposure, fish produced fewer eggs, but egg size was no different than those of
normoxia-exposed fish. This finding is consistent with reports that hypoxia exposed fish
have significantly reduced fecundity (Landry et al, 2007).
The hypoxia-induced
decrease in fecundity has been attributed to the disruption of normal testosterone and
estradiol levels in the female fish after chronic exposure to hypoxic conditions (Landry et
al, 2007; Wu et al, 2001).
However, the mechanisms underlying the decreased
fecundity in this study were not assessed.
Egg Components Volume
Interestingly, the relative decrease in whole egg volume of eggs produced by
hypoxia-exposed zebrafish was not accompanied by a reduction in yolk volume
(Fig.5.7). This caused the perivitelline volume to be significantly smaller in these eggs.
The function of the perivitelline space, the cavity between the outer surface of the yolk
134
and the inner surface of the chorion, includes protection of the embryo, nutritive supply,
and arguably, the regulation of embryonic metabolism and gas exchange (Laale 1980;
Burggren, 1985; Seymour, 1995). However, the direct role of the thickness or volume of
the perivitelline space in fish embryonic development remains largely unexplored.
Results of the present study suggested that parental hypoxia exposure induced
significant changes in egg component volumes, which may play a role in the
subsequent regulation of hypoxia resistance of the zebrafish larvae.
However this
remains highly speculative since no studies are known to have experimentally altered
perivitelline space assessed hypoxia resistance in zebrafish.
Summary
Collectively, the findings of the present study suggest that parental exposure to
hypoxia can induce a beneficial (although somewhat short-lived and age-dependent)
change in the hypoxia resistance of individuals of the subsequent generation, but that
this phenomenon is dependent on the amount of time the parents are exposed to
hypoxia. These studies extend and expand upon the current literature pertaining to
advantageous, environmentally-induced parental effects.
The finding that hypoxia-
induced parental effects modulate larval hypoxic stress response, but not thermal stress
response, of the next generation highlight the need for better understanding of the
mechanisms that underlie this phenomenon.
135
B. 2 weeks
Time to Loss of Equilibrium (seconds)
A. 1 week
120
120
100
100
80
80
60
60
40
40
20
20
0
0
0 4
6
8
10
12
14
16
18
0 4
20
C. 3 weeks
120
120
100
100
80
80
60
60
40
40
20
20
6
8
10
12
14
16
18
20
10
12
14
16
18
20
D. 4 weeks
0
0
0 4
6
8
10
12
14
16
18
20
0 4
6
8
Days Post Fertilization
FIGURE 5.1 Hypoxia resistance of larval offspring of zebrafish exposed to hypoxia
(square, broken line) or normoxia (circle, solid line) for 1 (A), 2 (B), 3 (C) or 4 (D) weeks.
A box indicates no significant difference among the groups. N’s > 16 for each group at
each time points. Data are presented as mean + S.E.M.
136
B.
Time to Loss of Equilibrium (seconds)
A.
120
120
100
100
80
80
60
60
40
40
20
20
0
0
0 4
120
6
8
10
12
14
16
18
0 4
20
C.
120
100
100
80
80
60
60
40
40
20
20
6
8
10
12
14
16
18
20
6
8
10
12
14
16
18
20
D.
0
0
0 4
6
8
10
12
14
16
18
20
0 4
Days Post Fertilization
FIGURE 5.2 Hypoxia resistance of larval offspring of zebrafish exposed to hypoxia
(square, broken line) or normoxia (circle, solid line) for 1 (A), 2 (B), 3 (C) or 4 (D) weeks.
Each plot within a graph represents a single clutch. A box indicates no significant
difference among the groups. N’s > 8 for each group at each time points. Mean +
S.E.M is presented.
137
CTMax or CTMin (Celsius)
42
41
40
39
12
11
10
9
0
10
20
30
40
50
60
70
Time Post-Fertilization (days)
Fig. 5.3 Critical thermal limits of zebrafish larvae of parents exposed to normoxia
(circle, solid line) or hypoxia (square, broken line) for 12 weeks. CTMax and CTMin
were not significantly different between treatment groups (two-way ANOVA with Tukey‟s
post hoc analyses, p > 0.05). N’s > 7 for each group at each time points. Mean +
S.E.M is presented.
138
Standard Body Length (mm)
6.0
6.0
A. 1 week
5.5
5.5
5.0
5.0
4.5
4.5
4.0
4.0
0.0
0.0
04
6.0
B. 2 weeks
6
8
10
12
14
16
18
04
20
6.0
C. 3 weeks
5.5
5.5
5.0
5.0
4.5
4.5
4.0
4.0
6
8
10
12
14
16
18
20
10
12
14
16
18
20
D. 4 weeks
0.0
0.0
0 4
6
8
10
12
14
16
18
20
0 4
6
8
Days Post Fertilization
FIGURE 5.4 Mean body length of larval offspring of zebrafish exposed to hypoxia
(square, broken line) or normoxia (circle, solid line) for 1 (A), 2 (B), 3 (C) or 4 (D) weeks.
A box indicates no significant difference among the groups. N’s > 16 for each group at
each time points. Mean + S.E.M is presented
139
Standard Body Length (mm)
6.0
6.0
A.
5.5
5.5
5.0
5.0
4.5
4.5
4.0
4.0
0.0
0.0
04
6.0
B.
6
8
10
12
14
16
18
20
0 4
6.0
C.
5.5
5.5
5.0
5.0
4.5
4.5
4.0
4.0
6
8
10
12
14
16
18
20
6
8
10
12
14
16
18
20
D.
0.0
0.0
04
6
8
10
12
14
16
18
20
0 4
Days Post Fertilization
FIGURE 5.5 Mean body length of larval offspring of zebrafish exposed to hypoxia
(square, broken line) or normoxia (circle, solid line) for 1 (A), 2 (B), 3 (C) or 4 (D) weeks.
Each plot within a graph represents a single clutch. A box indicates no significant
difference among the groups. N’s > 6 for each group at each time points. Mean +
S.E.M is presented
140
Fecundity (# of eggs clutch -1)
700
(4)
600
(4)
500
400
(3)
300
(3)
200
(4)
(3)
100
(4)
(3)
0
0
2
4
6
8
10
12
Parental Exposure Time (Weeks)
FIGURE 5.6 Fecundity of adult zebrafish exposed to normoxia (circle, solid line) or
hypoxia (square, broken line) for 1, 2, 6 and 12 weeks. A box indicates no significant
difference among the groups. N values for each mean value are indicated in
parentheses.. Mean + S.E.M is presented
141
Egg Parameters Volume (mm 3)
1.2
Whole Egg
1.1
1.0
Perivitelline
0.9
0.8
Yolk
0.2
0.1
0.0
0
2
4
6
8
10
12
Parental Exposure Time (Weeks)
FIGURE 5.7 Egg and egg component volumes of eggs of adult zebrafish exposed to
normoxia (unfilled symbol, solid line) or hypoxia (filled symbol, broken line) for 1, 2, 6 or
12 weeks. A box indicates no significant difference among the groups. N’s > 90 for
each group at each time points. Mean + S.E.M is presented
142
CHAPTER 6
PARENTAL EFFECTS DURING EARLY DEVELOPMENT OF VERTEBRATES:
CONCLUSIONS AND FUTURE DIRECTIONS
Maternal Effects of Yolk Hormones
An animal‟s phenotype is determined by its genotype as well as its environment.
During early embryonic development, the environment of the embryo is largely defined
by the current maternal condition.
In oviparious species, the egg environment
comprises the egg-shell, the albumen and the yolk.
The yolk, especially, contains
maternally-derived hormones, thyroxine (T 4), triiodothyronine (T3) and testosterone
(TE), that have the potential to dramatically influence early development when the
embryo‟s hormonal systems are not yet fully functional. In Chapter 2, the chronotropic
effects of perturbations in yolk hormones early in development were investgated in the
developing chicken embryo.
It was demonstrated that exogenous T 3 induced both
chronic and acute chronotropic effects, T 4 had only chronic effects, and TE had minimal
effect on heart rate during the first 4 days of development. Moreover, T 4 affected heart
rate in an age-dependent manner, with younger embryos (~64 h old) showing relative
tachycardia, while older embryos (~96 h old) showing relative bradycardia.
The
chronotropic cardiac effects were not accompanied by any hormone-induced changes in
developmental rate, suggesting that these effects were not due to increased maturity of
the embryo. These results highlight the complexity of the early embryo, as well as the
strong potential for maternal control of early developmental patterns, and possibly alter
the developmental trajectory of the animal.
143
Further extending the investigation of maternal effects of egg yolk in early
development, Chapter 3 characterized the role of inter-species variation in yolk
hormone concentrations in the determination of species-specific embryonic phenotype.
It was demonstrated that inter-species variation in egg size was a significant predictor of
yolk T3 and yolk TE in the egg, and that yolk T 3 and yolk TE accounted for the variation
in length of prenatal development among precocial avian species. In the second part of
the study, chicken embryos served as the model by which to assess the physiological
significance of the relationships among yolk hormones, egg mass and length of
incubation period among species. Removing chicken embryos from their original yolk
and culturing them on the yolks of quail, duck, turkey, goose, emu and ostrich yolk
resulted in significant changes in heart rate, mass-specific
manipulated embryo.
and body mass of the
Moreover, the variation in these traits were significantly
accounted for by the inter-species variation in yolk T3, yolk TE, incubation period and
egg mass of the species from which the yolk was derived. This novel finding strongly
suggested that maternal investment of yolk factors occurred in a species-specific and
taxon-specific manner which related to the egg size and incubation period characteristic
of the species. Although yolk hormones, egg mass and incubation period serve as
significant predictors of embryonic heart rate, metabolic rate and body mass, the
percentage of the variation accounted for was relatively small (~4 – 28%). This opens
up the door for investigation of other maternally-derived factors that contribute to the
variation induced by changes in yolk environment during early embryonic development.
In Chapter 4, the inter-species phenomenon in Chapter 3 was investigated on an
intra-species level. The domesticated chicken has long been artificially selected for
144
certain features such as high egg yield (layers) or large body mass (broilers). The
ability to harvest layer and broiler chicken embryos from their native yolk and expose
them to the yolks of other chicken breeds allowed for the elucidation of the extent to
which the breed-specific phenotype of early chicken embryos are determined by
maternally-derived yolk factors. It was demonstrated that exposure of layer and broiler
embryos on the yolk of another breed caused the embryos to adopt the phenotype of
the breed which represents that foreign yolk. However, the effect of yolk exchange on
embryonic heart rate, mass-specific oxygen consumption and body mass was largely
dependent on the breed of the embryo. This reflects the complexity of the interaction
between yolk environment and embryo genotype. Subsequent analyses of the yolk T 3
and yolk TE of the eggs of broilers and layers revealed that, although there were
significant differences in yolk hormones between the two breeds, it is unlikely that these
differences were responsible for the yolk-induced changes in physiology and
morphology observed.
For these studies (Chapter 2, 3 and 4), the mechanisms that underlie the
physiological changes caused by alterations in the yolk environment should ultimately
be investigated.
An assessment of hormone receptor density of cultured embryos
would be useful in understanding if yolk-induced changes in physiology and morphology
involve cellular changes that may potentially alter the trajectory of embryonic
development. Also, the comparison of proteomic profiles of the embryos cultured on the
yolks of different species and breeds, via molecular techniques such as 2-dimensional
gel electrophoresis, can lead to a better understanding of how yolk environment
influence the physiology of the early-stage embryo.
145
Hypoxia-Induced Parental Effects
The last study (Chapter 5) investigated parental effects from an inter-populational
perspective to determine whether the parental environment, rather than phylogeny, can
influence the physiology of the animal.
In zebrafish, parental exposure to hypoxia
increased the hypoxia resistance and body lengths of the larval offspring compared to
those from normoxia-exposed parents. However, the thermal tolerance of zebrafish
larvae was not affected by the environmental condition of the parents. Collectively,
these findings indicate that a change in oxygen availability in the parental generation
can induce a beneficial change in the subsequent generation, which is specific to the
parental stressor. An assessment of egg parameters of the hypoxia and normoxiaexposed fish revealed that hypoxia-exposed adults produced fewer eggs per clutch and
smaller eggs with smaller perivitelline volume. The change in perivitelline volume is
suggestive of an egg-induced mechanism for the heightened hypoxia resistance
observed in the offspring of hypoxia exposed zebrafish. However it is not known if
perivitelline
volume
can
affect
zebrafish
physiology
and
morphology
during
development. A fruitful direction of investigation would be to assess the difference in
yolk factors (i.e. hormones and mRNAs) between the two experimental groups. Also, a
comparison of the protein profiles of the larvae of hypoxia-exposed and normoxiaexposed fish would shed light on the mechanisms by which hypoxia resistance is
altered between these two groups.
In summary, parental effects was important in modulating the morphology and
physiology of chicken embryos and zebrafish larvae, and thus may have a role in
altering the developmental trajectory of the animal.
146
Future studies should employ
biochemical and molecular approaches to further elucidate the mechanisms by which
parental effects influence early developing systems.
147
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