University of Southern California Dissertations and Theses

Skin appendage growth control by hormones and morphogens

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Skin appendage growth control by hormones and morphogens

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SKIN APPENDAGE GROWTH CONTROL BY HORMONES AND
MORPHOGENS

by

Julie Ann Mayer

____________________________________________________________________

A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(PATHOBIOLOGY)

December 2007

Copyright 2007 Julie Ann Mayer

ii
TABLE OF CONTENTS

List of Tables.......................................................................................................... iv
List of Figures.......................................................................................................... v
Abstract ................................................................................................................. vii

Chapter 1: Introduction ............................................................................................ 1
1.1 Sex hormone pathways ............................................................................. 3
1.2 Sexual dimorphism in mammals ............................................................... 6
1.3 Sexual dimorphism in birds ...................................................................... 8
1.4 Problems related to sex hormone-dependent skin appendage
morphogenesis: alopecia, hirsutism, and trans-differentiation ................. 11
1.5 Sex hormone-dependent genetic diseases................................................ 14
1.6 Sex hormone-dependent tumor growth ................................................... 16
1.7 The need for establishing an in vivo experimental model for sex
hormone-dependent growth .................................................................... 20
Chapter 2 : Hormone regulation of skin appendages: Rooster and hen tail feathers. 22
2.1 Classical Experiments............................................................................. 22
2.2 Effects of Ovariotomy on hens................................................................ 23
2.3 Methods and Materials ........................................................................... 24
2.3.1 Housing of birds ............................................................................. 24
2.3.2 Measurement of feather growth....................................................... 24
2.3.3 Sections and immunostaining.......................................................... 25
2.3.4 In-situ hybridization........................................................................ 25
2.3.5 RT-PCR.......................................................................................... 26
2.3.6 ELISA ............................................................................................ 26
2.3.7 Hormone and Inhibitor Injections.................................................... 26
2.3.8 Hormone inhibition by implanting pellets ....................................... 27
2.3.9 Microarray...................................................................................... 27
2.4 Results.................................................................................................... 28
2.4.1 Different growth kinetics in male- and female-type tail feathers...... 28
2.4.2 Comparison of feather follicles in male and female rectrice feathers 32
2.4.3 Comparison of feather branches rectrice feathers ............................ 37
2.4.4 Determining the effects of hormone modulation by local injection.. 42
2.4.5 Hormone pathway inhibition studies ............................................... 48
2.4.6 Local conversion of hormones ensures a female phenotype............. 54
2.5 Discussion .............................................................................................. 57
2.5.1 Sex hormones in epithelial–mesenchymal interactions .................... 57
2.5.2 Potential cellular/molecular mechanisms......................................... 62
2.5.3 Regional specificity of sex hormone effects .................................... 66

iii
2.6 Summary ........................................................................................ 68
Chapter 3 : Mouse Mammary gland project............................................................ 71
3.1 Introduction............................................................................................ 71
3.2 Methods and Materials ........................................................................... 73
3.2.1 Production/Genotyping of the transgenic KRT14-Noggin mouse .... 73
3.2.2 Histological, histochemical, and Immunohistological Staining ........ 74
3.2.3 In-situ hybridization........................................................................ 75
3.2.4 Induction of involution.................................................................... 76
3.3 Results.................................................................................................... 76
3.3.1 Mammary gland development in KRT14-Noggin mice ................... 76
3.3.2 BMP4 is increased in the mesenchyme of KRT14-Noggin
involuting mammary glands............................................................ 82
3.3.3 Delayed mammary gland involution in KRT14-Noggin mice.......... 84
3.3.4 Collagen I is maintained in the mammary glands of
KRT14-Noggin mice after involution.............................................. 87
3.4 Discussion .............................................................................................. 90
3.4.1 The loss of the BMP pathway via Noggin overexpression delays
mammary gland involution while increasing the amount of
collagen I present............................................................................ 90
3.4.2 BMP and Human Disease (Cancer) ................................................. 91
Chapter 4 : Mouse nipple ....................................................................................... 93
4.1 Introduction............................................................................................ 93
4.2 Methods and Materials ........................................................................... 97
4.2.1 Animals .......................................................................................... 97
4.2.2 Histological Histochemical and Immunohistological Staining ......... 97
4.2.3 In situ hybridization ........................................................................ 98
4.2.4 Scanning Electron Microscopy........................................................ 98
4.2.5 Semi Quantitative RT-PCR............................................................. 99
4.3 Results.................................................................................................. 100
4.3.1 Overexpression of Noggin leads to the conversion of nipple
epithelium into competent hair follicle forming epithelium ........... 100
4.3.2 Overexpression of Noggin leads to ectopic expression of Shh
while suppressing PTHrP.............................................................. 104
4.3.3 Overexpression of Noggin leads to pigmented nipples .................. 106
4.4 Discussion ............................................................................................ 111
4.4.1 Signaling pathways involved in hair follicle and nipple formation. 111
4.4.2 Evolution of the nipple and mammary gland ................................. 117
Chapter 5 – Discussion......................................................................................... 118
Bibliography........................................................................................................ 122

iv
LIST OF TABLES

1. Tail feather physical characteristics—male versus female........................... 42
2. List of candidate genes generated from microarray analysis........................ 53
3. RT-PCR primers......................................................................................... 54

v
LIST OF FIGURES

1. Male and female adult chickens.................................................................... 2
2. Hormone biosynthetic pathway .................................................................... 5
3. How does the growth of male tail feathers differ from female tail feathers? 30
4. Growth kinetics of male and female tail feathers......................................... 31
5. Schematic of feather follicle structure......................................................... 33
6. Molecular expression in male and female tail feathers ................................ 36
7. Morphological difference in male and female tail feathers .......................... 40
8. DHT ELISA............................................................................................... 43
9. Phenotype of male and female birds injected with Flutamide or DHT......... 45
10. DHT induced alopecia in birds ................................................................... 47
11. Percentage of feathers remaining in anagen ................................................ 51
12. RT-PCR results for male and female feathers compartments....................... 55
13. Possible models of hormone effects............................................................ 59
14. Sex hormones' influence on feather diameter and length............................. 63
15. Whole mount Carmine staining of mammary glands................................... 78
16. Stromal increase and milk products sustained in the ducts .......................... 79
17. Immunostaining of wild-type and KRT14-Noggin mice .............................. 81
18. In-situ hybridization of the mammary glands.............................................. 83
19. H&E staining of wild-type and KRT14-Noggin involution MGs ................. 86
20. Trichrome staining of wild-type and KRT14-Noggin mammary glands ....... 88

vi

21. Collagen I staining of wild-type and KRT14-Noggin mammary glands ....... 89
22. Scanning electron microscopy and quantitation of hair follicles................ 101
23. Histological and immunohistochemical analysis of adult mouse nipples 104
24. Expression patterns for PTHrP, Shh, and pSmad1/5/8............................... 106
25. Nipple Pigmentation................................................................................. 108
26. Expression of the melanogenesis regulators in nipples.............................. 111
27. Effects of BMP signaling on epithelial appendages................................... 117

vii
ABSTRACT

The mammary gland, hairs, feathers, etc. are all skin appendages derived
from epidermis as a result of epithelial - mesenchymal interactions. The growth of
these epithelial organs shares fundamental morphogens including BMP, SHH,
Wnt/beta catenin, etc. Mutation or de-regulation of these morphogens have been
identified in many human diseases including tumors and genetic diseases. Sex
hormones also play a major role in regulating the growth and phenotypes of these
epithelial organs (sexual dimorphism). How the sex hormones are coupled to the
morphogens is mostly unknown. In this thesis I utilized two distinct epithelial
appendage models, a chicken tail feather (between rooster and hen), and the mouse
mammary gland model. In the chicken tail feather model, we were able to show that
the female feather phenotype is dependent on the local conversion of estrogen. We
also utilized the transgenic KRT14-Noggin (BMP antagonist) mouse that shows
abnormal mammary gland stroma formation and delayed involution. Imbalance of
BMP pathway lead to increased collagen I deposition surrounding the ducts and hair
formation within the nipple of these transgenic mice. Taken together, this opens the
possibility that the sex hormone pathway may be involved with the BMP pathway.
We offer novel experimental models that may contribute to further understanding of
the hormone dependent growth of breast and prostate cancer, which are clinically
more important but less susceptible to experimental analyses.

1

1.0 Chapter 1: Introduction
Sexual dimorphism is the systematic difference in phenotype between individuals of
different sex in the same species (Fig. 1). Sexual dimorphisms readily apparent on an
animal's body surface serve to attract members of the opposite sex. Frequently these
secondary sexual characteristics involve the size, shape, and color of epithelial
appendages, forming the basis for sexual selection (Darwin, 1871). The formation of
internal and external reproductive organs including the prostate and mammary
glands are essential for reproduction. The morphogenesis of these different
epithelium-derived organs results from epithelial–mesenchymal interactions and are
variations on top of a common theme (Chuong, 1998; Widelitz and Chuong, 1999;
Widelitz et al., 2003). One of the major variations is the physiological response
depending on the concentration of sex steroid levels, their receptors, and their co-
activators. Here we will first survey sex hormone pathways and its roles in the
morphogenesis of various epithelial organs in a variety of physiological and
pathological conditions. We will then present some pilot data toward establishing
chicken tail feathers as a model for the sex hormone-regulated organ formation in the
context of system biology/pathology. In the discussion, we hypothesize some
mechanisms through which sex hormone pathways may interface with
morphogenesis-related pathways.
2

Fig. 1 Male and female adult chickens. (A) Photo of a male white leghorn. (B) Photo
of a female white leghorn. (C) Left: male tail feather. Right: female tail feather. Both
are from the midline.
3

1.1 Sex hormone pathways
To obtain a mechanistic understanding of sexual dimorphism, we first have to
understand their biochemical/molecular pathways. The sex steroid biosynthetic
pathway is well established (reviewed in Chang, 2002). A simplified biosynthesis
pathway is briefly described here and shown schematically in Figure 2. Steroids are
synthesized from cholesterol, which is converted to pregnenolone by the side chain
cleavage enzyme, p450scc, within the mitochondria. The synthesis pathways
bifurcate to form progesterone and the corticosteroids or to form androgens and
estrogens. Progesterone can also be converted toward androgens using similar
enzymes as described below in a parallel pathway. Pregnenolone undergoes 17-α-
hydroxylation by microsomal P450c17. 17-hydroxy pregnenolone is converted to
dehydroepiandrosterone (DHEA) by the 17–20-lyase activity of P450c17. DHEA is
then converted to androstenedione by 3-β-hydroxysteroid dehydrogenase.
Androstenedione can be converted in a reversible reaction to testosterone by 17-β
hydroxysteroid dehydrogenase. Testosterone can then be converted to 5α
dihydrotestosterone (DHT) by the microsomal NADPH-dependent enzyme 5α-
reductase. The non-aromatic androgens, testosterone or androstenedione, can also be
converted by P450 aromatase to the aromatic estrogens, estradiol or estrone,
respectively. Estrone can also be converted to estradiol inareversible reaction by 17-
β hydroxysteroid dehydrogenase. Hence, in the biosynthetic pathway, androgens are
4

synthesized first and subsequently converted to estrogens in what is thought to be the
rate-limiting step in estradiol biosynthesis (Jarman et al., 1998).

In order for the steroid hormones to exert their effects, they must bind to specific
nuclear receptors, which trigger transcriptional activation events. Thus, sex
hormone-dependent pathways involve ligands, receptors, enzymes, and co-activators
(Chang, 2002). Along the pathways, different members play pivotal roles, and errors
in them can have serious patho-physiological consequences. To deal with the
consequences of these errors, drugs have been developed to target different steps
along these pathways.
5

Fig. 2 Hormone biosynthetic pathway. Enzyme names are indicated in green.
Products are indicated in black.
6

1.2 Sexual dimorphism in mammals
The roles of sex hormones on skin appendage dimorphisms have been documented
in a few species. Both hormones (androgens and estrogens) are involved in
determining the type of hair produced by follicles in mammals (Randall, 1994). Here
we introduce some concepts of hormone influence on skin appendage sexual
dimorphisms.

Primates: Pelage and skin can be markedly dimorphic in primates. The brightly
colored face of male mandrills as compared with that of females is one of the most
well-characterized examples (Setchell and Dixson, 2001). Male gelada baboons also
exhibit skin and pelage differences. They have a large prominent mane, which
probably serves to make males appear larger (Crook, 1972). Male orangutans possess
large cheek flanges and laryngeal pouches. Many species of male and female
primates differ dramatically in pelage color, while others show minor sex differences
in pelage color or patterning (Crook, 1972; Hershkovitz, 1977). The reasons for these
pelage and skin dimorphisms remain unclear.

Men and women differ in the amount of hair developed and the places at which hair
will develop. Androgens paradoxically can stimulate and inhibit human hair growth
in different regions or at different times. The hair follicle is a complicated endocrine
target organ because its response to androgens is extremely complex and this
paradox is unique in endocrinology (Randall et al., 2000). Low-capacity, high-
7

affinity androgen receptors (ARs) are found in all dermal papilla cells from
androgen-sensitive sites (Randall et al., 2000). Even within these androgen-sensitive
hair follicles there are differences. Androgens can stimulate the beard and body hair
growth while inhibiting scalp hair growth. Presumably, this is due to differential
gene expression in the hair follicles of these different regions caused during
development (Randall et al., 2000). In women, the skin is a major site of testosterone
formation. Half of their testosterone production stems from the conversion of
secreted 17-ketosteroids including DHEA and androstenedione (Rosenfield and
Lucky, 1993), whereas in men, the testes are the major site of testosterone
production. In humans, very coarse, pigmented, sexual hairs develop during puberty
from fine, unpigmented, vellus hairs. Genetic susceptibility in certain individuals
allows for androgens to act upon the terminal hair follicles causing them to become
vellus ones leading to androgenic alopecia (Randall et al., 2000).

Deer: In red deer sexual dimorphism is displayed in terms of size, with the males
being much larger in size than the females (Post et al., 1999). Mates are selected
based on strategies of growth in relation to reproduction. The red deer exhibits
highly coordinated seasonal coat growth with two distinct pelage types each year.
During the winter breeding season the male red deer grows an androgen-dependent
mane due to an increase in levels of plasma testosterone (Lincoln, 1971; Thornton et
al., 2000). This is then replaced with short hairs in the summer when the levels of
plasma testosterone drop. Antlers are derived from the bone but also exhibit
8

hormone-dependent growth. The hairs on newly formed skin over the antler and the
mane of the deer (Thornton et al., 2001) can also be used as a model to study sex
hormone-dependent hair growth.

Mice: In mice, sexual dimorphisms are not clearly visible. Both androgens and
estrogens have been implicated in regulating the hair cycles. Estrogen receptors
(ERs) have been located in the dermal papillae of telogen follicles (Oh and Smart,
1996) and estradiol has a direct inhibitory effect on the hair fiber growth cycle
(Mohn, 1958; Ebling and Johnson, 1964; Oh and Smart, 1996). It has important to
note that it is ER α and not ER β that has been localized in the hair follicles of mice
(Chanda et al., 2000). It is also been found that pituitary prolactin regulates seasonal
hair follicle growth cycles in many mammals as well as in the non-seasonal
laboratory mice pelage replacement (Craven et al., 2001). Prolactin appears to play
an inhibitory role in the pelage replacement of many mammals.

1.3 Sexual dimorphism in birds
Many birds show sexual dimorphisms. Mainly, it is the male that exhibits fancy
feathers or other integument appendages (e.g., comb, wattle) to attract the female.
Feathers come in different sizes, shapes, colors, etc. Contour feathers are found on
the body while rectrice feathers are found on the tail. The longest feathers are the tail
feathers of the peacock and those of birds of paradise. While these traits make

9

individuals spectacularly beautiful and increase their chance for mating, it also
increases their visibility and hence decreases their survival due to predation. This
observation puzzled Darwin in his contemplation of sexual dimorphism (Darwin,
1871).

In chickens, male and female chicks are the same weights upon hatching and do not
exhibit any distinguishing secondary sexual characteristics with the exception of
specific genetic variants in down color or wing feather length (Jacob and Mather,
2000). The male will develop a larger body, comb, and wattles than the female. The
male white leghorn's comb is turgid and stands erect; whereas the female's comb
upon full maturity is less turgid and lies over to one side. The male also develops a
larger spur and has longer and more pointed hackle feathers than the female (Jacob
and Mather, 2000).

It is widely known that male tail feathers grow longer and wider than female tail
feathers and have different shapes (Aparicio et al., 2003). Here, size is measured as
the length of the feather and shape is measured as the varying width of the vane and
the curvature of the rachis. This difference has been attributed to a benefit to male
birds with longer tails during the sexual selection process. Tail length has been
negatively correlated with the density of barbs and rachis width in males but not in
females (Aparicio et al., 2003). Testis size has also been positively correlated with
dimorphisms in wing and tail length (Dunn et al., 2001). Hence, these attributes
10

appear to be sex dependent and thus hormone regulated. George and Wilson (1980)
suggested that female feathering patterns in chickens result from ovarian estrogens,
which leads us to believe that male feathering patterns result from plasma
testosterone levels. We believe that the feather offers an opportunity to examine sex
hormone effects on epithelial–mesenchymal interactions since the difference in
feather length and diameter is sex dependent and thus hormone regulated. This
hypothesis is further supported by the following observations.

Since male birds can be aggressive, small chicken farms will sometimes castrate
their roosters at 2–4 weeks of age. This blocks androgen synthesis, making the birds
docile with tender meat. The combs and wattles diminish in size and resemble those
of female birds, but the hormone-sensitive feathers (i.e., tail feathers) grow longer.
These birds are called capons (Payne, 1936; Jacob and Mather, 2000). Capons with
similar feathering patterns previously were made by administering stilbestrol (an
estrogen) rather than performing castrations. Caponization later in life produces a
female feathering pattern. Nowadays, a combination of changes brought about by
breeding and diet produces capons that mature more rapidly than in the past. Ten
pound birds can be raised in 12–13 weeks as opposed to 20 weeks in years past. The
feathers of 10 pound capons no longer grow as long as they used to (they mature
prematurely).

11

On the other hand, the left ovary of female birds is functional while the right ovary
regresses under natural conditions (Andrews et al., 1997). Upon ovarectomy, the
right gonad can masculinize and produce androgen (Groenendijk-Huijbers, 1965),
resulting in a "male" feathering pattern (Frankenhuis and Kappert, 1980). It is
unknown whether this is due to a loss of estrogen or a gain of androgen.

1.4 Problems related to sex hormone-dependent skin appendage
morphogenesis: alopecia, hirsutism, and trans-differentiation
In humans and other animals, the growth of skin appendages can go wrong due to
errors along the sex hormone pathway. The pathogenesis may be at the level of
hormone concentration, receptors, enzymes, or co-activators/co-suppressors. The
density and size of skin appendages may increase or decrease. Alopecia can occur in
males or females (Randall, 1994; Birch et al., 2002) and likely follow different
etiologies. Dermatologists are trying to identify causes of alopecia so that these
conditions can be corrected, perhaps by relieving a pathological blockade.

Androgenic alopecia: Commonly referred to as "male pattern baldness", androgenic
alopecia is also the most common form of hair loss in women. Androgenic alopecia
is the name for male and female pattern baldness. This androgenetic alopecia is
present in at least 50% of males by age 50, 70% of males in later life, and 30–40% of
females over 70 years of age (Norwood, 2001). Androgenetic alopecia is

12

characterized by the progressive replacement of terminal hairs with vellus hairs
(Paus and Cotsarelis, 1999; Hoffmann and Happle, 2000). This is fostered by the
continually shortening anagen phase of the hair cycle causing a higher percentage of
telogen phase hairs in the crown and forehead region. Furthermore, the time between
telogen and a new anagen phase increases, leading to reduced hair numbers (Courtois
et al., 1994). Another intriguing observation is the regional-specific response of hair
growth to androgen. This was found to be partially due to the differential expression
of 5α-reductase type 2 in the dermal papillae of scalp hair follicles (Thigpen et al.,
1993). Inhibition of this isozyme with finasteride was shown to be significantly
effective in the treatment of androgenetic alopecia (Whiting, 2001; Shapiro and
Kaufman, 2003).

Females can also suffer from hirsutism or alopecia. Female-type alopecia due to
hyperandrogenism may be due to androgen metabolism, whereas this pathway plays
a very minor role, if any, in postmenopausal women suffering from female-type
alopecia (Shum et al., 2002). Treatment of women suffering from idiopathic
hirsutism with anti-androgen drugs reduces the size of body and facial hair by
reducing the formation of a medulla (Ebling, 1987). Still, how DHT promotes cell
proliferation or suppresses hair differentiation in different hair follicles (scalp hairs
versus beards versus axilla hairs) remains unknown. The interactions are further
complicated by the interactions of circulating steroids and the sex hormone
processing enzymes in different follicles (Hoffmann and Happle, 2000).
13

Our nearest non-human primate relatives, orangutans and gorillas, can also develop
androgenetic alopecia. The stump tailed macque has been used as an in vivo model
for androgenic alopecia (Brigham et al., 1988). Once hair follicles have been
exposed to androgens they are fated to become androgen sensitive and androgenetic
alopecia can develop. Androgenetic alopecia develops as a gradual reduction of scalp
hair follicle size, accompanied by reduced time in the anagen active growth phase,
leading to more hair follicles in the telogen resting stage of the hair cycle. Although
periods of anagen are reduced, catagen and telogen time periods remain the same. In
hair, the hormone acts first on the DP, which then signals to and induces growth in
the epithelium (Obana et al., 1997; Randall et al., 2001).

In chickens, a dramatic example of hormone-dependent growth is the conversion of
male into female feather phenotypes. In "henny feathering", a genetically transmitted
constitutively active aromatase in the skin can cause roosters to exhibit "female-
type" tail feathers (Wilson et al., 1987). Male chickens carrying the henny feathering
trait virilize normally but develop a female feathering pattern (George and Wilson,
1980). This autosomal dominant mutation causes the accumulation of aromatase
mRNA and activity in extragonadal chicken tissues (Matsumine et al., 1991), which
converts androgen to estrogen in the skin. Again, it is unknown if this is due to a
decrease of androgen or an increase of estrogen.
14

1.5 Sex hormone-dependent genetic diseases
The development of urogenital organs and external genitalia is essential to carry out
reproductive function. These epithelial and mesenchymal tissues are malleable and
can form the male or female type during embryonic development. As a result,
patients who suffer from inborn errors of the sex hormone pathway may produce
epithelial organs of the wrong sexual type.

5α-reductase deficiency: There are two forms of 5α-reductase that can convert
testosterone to DHT. They are differentially expressed in various tissues. Androgen
action in sexual organs is primarily dependent upon the type 2 isozyme (Thigpen et
al., 1993) and deficiency of this isozyme form leads to pseudohermaphroditism
(Andersson et al., 1991). There is only one wave of expression of the type 2 isozyme
that starts at birth and ends by 3 years of age. The type 2 isozyme is not detected in
adult skin but is found in the hair follicles of the scalp, suggesting that balding may
be pre-determined early in development (Bayne et al., 1999). The major form of 5α-
reductase in the skin is the type 1 isozyme, which is expressed in two waves. The
first occurs at birth and lasts until 3 years of age and the second begins during
puberty and lasts throughout life (Thigpen et al., 1993).

15

Patients with 5α-reductase deficiency fail to metabolize testosterone into DHT.
Defects in 5α-reductase typically result in an intersex phenotype. Intersexed
individuals do not develop pubic, axilla, or beard hairs normally (Griffin and Wilson,
1989), but they do exhibit normal scalp hair development (Randall et al., 1991). This
suggests that the conversion of testosterone to DHT by 5α-reductase is not essential
in follicles that are androgen sensitive in both sexes but only in those that distinguish
the adult male (Randall et al., 2000).

Pseudohermaphroditism: Male pseudohermaphroditism is caused by a defect in
testosterone biosynthesis. Female pseudohermaphroditism is typically caused as a
result of a defect in the enzymes leading to glucocorticoids or mineralocorticoids
causing a shunting of precursor molecules into the androgenic pathway (Haqq and
Donahoe, 1998) or as a result of defects in estrogen formation.

Androgen insensitivity syndrome: Androgen insensitivity syndrome is also referred
to as testicular feminization, which is a subtype of male pseudohermaphroditism. A
defect in the AR leads to an overall resistance to androgens in this disorder. Males
with androgen insensitivity syndrome will typically have no hair growth in the
axillary and pubic regions (Hamilton, 1950). Expression of aromatase in genital skin
is androgen dependent, as demonstrated by lower levels of aromatase present in cells
from patients with androgen insensitivity syndromes (Stillman et al., 1991).

16

1.6 Sex hormone-dependent tumor growth
Cancer rates continue to rise here in the United States mainly due to increasing rates
for prostate cancer among men and for breast cancers among women (Devesa et al.,
1995) while mortality is steadily decreasing (Quinn, 2003). Some of these tumor
types are sex hormone dependent while some are not. For those with sex hormone
dependence, it is possible to treat the tumor by blocking sex hormone synthesis or
receptor binding.

It has been suggested that along with breast cancer, other cancers including ovarian
and endometrial cancers are a direct result of ovarian function (Pike et al., 2004).
Oophorectomy then would significantly reduce the incidence of all three of these
cancer types (Pike et al., 2004), but this would not be a proper solution for most
women. The incidence of breast, endometrial, and ovarian cancers increases with
age, but shows a notable slowdown in relation to age around the time of menopause
(Pike et al., 2004). Menopause clearly has a large protective effect against all three of
these cancers. One explanation for this is that ovarian hormones stimulate cell
division which is reduced after menopause. The ovaries begin producing less
steroids, in particular estrogen as menopause progresses and thus the relevant tissues
cease to divide as a result of lack of steroid hormones. This protective effect of
menopause and lack of steroid hormones remains the central clue to the
17

chemoprevention of various types of hormone-dependent cancers including breast,
endometrial, and ovarian cancers (Pike et al, 2004).

Estrogen and breast cancer: Breast cancer is the second leading cause of cancer-
related deaths among women in the United States. The American Cancer Society
estimates that 40,200 (39,800 females, 400 males) people will die from breast cancer
in 2003. Sex steroids have been suggested to be involved in breast carcinogenesis,
and may be fundamental molecules involved in the progression and growth of many
carcinomas. In many breast cancer types, tumorigenesis initially is hormone
dependent. Conditions associated with altered estrogen/androgen ratios predispose
patients to male breast cancer (Ravandi-Kashani and Hayes, 1998). There are many
risk factors associated with the onset of breast cancer. The three most important and
well-studied reproductive risk factors for breast cancer include early menarche, late
first full-term pregnancy, and late menopause (FFTP; Kelsey and Bernstein, 1996;
Colditz and Rosner, 2000).

The mammary gland is an epithelial organ derived from epidermis as a result of
epithelial–mesenchymal interactions (Veltmaat et al., 2003; Parmar and Cunha,
2004). Estrogens play an important role in the growth and differentiation of normal
breast epithelium acting through the ER in embryonic development and in lactation
cycle. Hormone therapy tries to modulate pathway activity at different levels.
Estrogen antagonists block estrogen action in several ways. For example,
18

tamoxifen competitively binds to the ER, displacing the natural ligand, estradiol.
Aryl hydrocarbon receptor (AhR) agonists down-regulate ligand-bound ER
(Wormke et al., 2003), but up-regulate unbound ER (Ohtake et al., 2003). Aromatase
inhibitors block the conversion of testosterone to estradiol (Elbrecht and Smith, 1992
). Aromatase is the enzyme responsible for the conversion of non-aromatic
androgens, particularly testosterone and androstenedione, to aromatic estrogens:
estrone and estradiol. In postmenopausal women, testosterone is converted to
estrogens by aromatase in the skin, muscles, and liver

Aromatase inhibitor in breast cancer therapy: The conversion of androgens to
estrogens by the enzyme aromatase is considered the rate-limiting step of estrogen
biosynthesis (Brodie and Njar, 2000). As women proceed through menopause,
ovarian estrogen production diminishes. In postmenopausal women, the main source
of estrogen comes from the conversion of androgens to estrogens by aromatase in
peripheral tissues such as adipose and muscle tissues (Brodie and Njar, 2000). Since
estrogens are known to play a critical role in the development and growth of breast
cancers, inhibitors of aromatase have been produced in an effort to inhibit tumor
proliferation. Aromatase inhibitors can be divided into two classes: steroidal and
non-steroidal inhibitors. Steroidal inhibitors such as formestane and exemestane,
compete with androgens (testosterone and androstenedione) where they bind
irreversibly to aromatase causing inhibition (Buzdar and Howell, 2001). Non-
steroidal inhibitors including fadrozole, anastrozole, and letrozole compete with
19

androgens for the conversion of testosterone to estradiol (Buzdar and Howell, 2001).
Androgenic alopecia is common in women undergoing hormone therapy as treatment
for breast cancer (Buzdar et al., 2001). Mutations in the aromatase gene can also
cause females to develop a form of pseudohermaphroditism with androgenetic
alopecia (Hoffmann and Happle, 2000).

Androgen and prostate cancer: Prostate cancer is the second leading cause of
cancer-related deaths among men in the United States. The American Cancer Society
estimates that 28,900 men will die from prostate cancer in 2003. It is estimated that
there is a one in six probability that a man will develop prostate cancer in his
lifetime. Prostate cells are dependent upon androgens (testosterone and DHT) for
growth and function (Anderson, 2003; Marker et al., 2003). Males who are castrated
at an early age do not develop prostate cancer. This suggests that androgens and their
receptors are major risk factors for prostate cancer.

Prostate cancers initially arise as androgen dependent but following androgen
ablation therapy, the tumors re-surface in an androgen-independent manner. Anti-
androgens can compete for AR binding sites in the nucleus of the prostate cells, thus
inhibiting prostate cancer growth and promoting apoptosis (Anderson, 2003). One of
the treatment modalities utilizes anti-androgens (flutamide, casodex) to suppress
androgen activity. Flutamide is a non-steroidal pure antiandrogen that acts as a
competitive inhibitor of DHT (Grzywacz et al., 2003) but does not suppress
20

testosterone secretion (Anderson, 2003). Hydroxyflutamide is the active metabolite
of Flutamide and blocks the expression of genes containing an androgen response
element in their promoters while preventing stabilization of the AR (Grzywacz et al.,
2003). Other more recent treatments utilize agents such as finestaride to inhibit
androgen synthesis.

The AR may serve as a link between other signaling pathways. β-catenin (Truica et
al., 2000) and Her2/neu (Zhau et al., 1992) have been reported to interact with the
AR in prostate cancer to elicit cell survival and growth. Members of the bone
morphogenetic protein (BMP) pathway have also been implicated in regulating
prostate cancer growth. BMP2 has been shown to decrease prostate cell growth (Ide
et al., 1997) and others such as BMP4 (Thomas et al., 1998) and BMP6 (Barnes et
al., 1995) are constitutively expressed despite castration.

1.7 The need for establishing an in vivo experimental model for sex hormone-
dependent growth
A novel, physiological epithelial growth model responsive to sex hormones: In order
to manage the loss of new growth in alopecia, an understanding of how nature
regulates normal new growth must be achieved. The mechanism of how sex
hormones regulate growth is largely unknown and model systems to examine it are
lacking. While mice have strong genetics, they are not good models because
they
21

lack sexual dimorphism. In humans, the hormone-regulated growth control of hair is
mostly studied in patients with abnormal hair formation. Thus, there is a need for a
good model in which both in vitro and in vivo studies can be performed to bring out
these mechanisms. We have been using White Leghorn chicken feathers as an animal
model to study sex hormone effects on epithelial appendage growth. Chickens have
clear cut sexual dimorphism (Yu et al., 2004) and the regulation of male and female
tail feathers is part of their physiological process. Roosters and hens are good
experimental models because they provide a true in vivo model with an obvious
sexual dimorphism in their tail feathers or "rectrices" (Fig. 1).

22

2.0 Chapter 2 – Hormone regulation of skin appendages: Rooster and hen tail
feathers
2.1 Classical Experiments
Feathers come in different sizes and shapes. Size is measured by the length. Shape is
measured by the varying width of the vane and the curvature of the rachis. Contour
feathers are found on the body and do not respond to sex hormones, whereas rectrice
feathers found on the tail are hormone responsive. Presumably, female chicken
feathering patterns result from ovarian estrogens and male feathering patterns from
androgens. However, there are several examples that raise paradoxes as to the roles
of steroid hormones in growth control. 1) Caponization, the castration of male birds
at 2-4 weeks of age, blocks androgen synthesis. The wattles and combs diminish in
size and resemble those of female birds, but the hormone sensitive feathers (ie. tail
feathers) grow especially long (Payne, 1936; Jacob and Mather, 2000). Historically,
capons were made by injecting stilbestrol (an estrogen) rather than by performing
castrations. Interestingly, if caponization is performed on the birds after hormones
are present it will produce a female feathering pattern. 2) Male golden Campine and
Sebright bantam chickens have an autosomal dominant mutation that causes them to
virilize normally in response to androgen stimulation but accumulate high levels of
Aromatase, which converts androgen to estrogen in the skin producing female
feathering characteristics (henny feathering; George and Wilson, 1980; George et al.,
1990; Matsumine, 1991). 3) Normally the left ovary of female birds is
functional
23

while the right ovary regresses (Andrews et al., 1997). Following ovarectomy, the
right gonad can masculinize and produce androgen (Woods et al., 1966) resulting in
a “male” feathering pattern (Frankenhuis and Kappert, 1980).

2.2 Effects of Ovariotomy on hens
Experiments involving the gonadectomy of birds have been performed in great
detail. In 1929, Domm and colleagues report on the phenotype of female hens having
underwent both single and bilateral ovariotomy (Domm 1929). Using brown leghorn
birds they show that complete bilateral ovariotomy leads to a neutral/asexual type
common to both sexes. This includes the display of very small head furnishings
(Comb and wattle) as compared to birds that underwent a sinistral (single left-side)
ovariotomy that display large male-like head furnishings. These birds that underwent
a sinistral ovariotomy also displayed male plumage that at a later period reverts to
the female type. A complete bilateral ovariotomy on the other hand, produced male
plumage that retained this character throughout the duration of the bird’s life. Spurs
developed in both sinistral and bilaterally ovarectomized birds and there were no
measurable differences in size, the birds remained the size of normal hens (Domm
1929).

In the above examples, it is clear that estrogens are responsible for creating the
phenotype of the female feather (Juhn 1930) perhaps by playing a major role on
24

the localized growth zones in tail feathers. The following studies will focus on the
tail feathers as the main model system, and utilizing current methodologies I set forth
to understand the molecular signaling that leads to the sexual dimorphism in birds.
Understanding how previous experiments show that estrogen is regulating the shape,
size and pigmentation of the female feather is of particular importance.
2.3 Methods and Materials
2.3.1 Housing of birds
The birds are housed within the USC Vivaria facility. They are monitored daily and
housed under a 12-hour light/dark cycle. The birds are fed lay crumble (Southwest
Farms, Perris, CA) and water ad lib.
2.3.2 Measurement of feather growth
Quantitative differences in size and shape were assessed using a regeneration model
because it allows for the synchronization of the feather growing process. Adult male
and female white leghorn chickens of known ages (SPAFAS, pathogen-free
chickens) were used in this experiment because they are sexually mature and should
possess ER, AR, and detectable levels of sex steroids. The tail feathers of these birds
were measured every other day for the duration of the regeneration cycle and
photographed periodically. For each bird, the two feathers closest to the midline
were measured since they have the most pronounced sexual dimorphic
phenotypes. The lateral feather to each side of the central pair was also
25

measured as a comparison

since they too contribute to the dimorphic phenotype. From the daily measurements,
the average rate of growth as well as total length and duration of growth were
quantified.
2.3.3 Sections and immunostaining
Specimens were fixed in 4% paraformaldehyde for 24 hr at 4°C followed by a
dehydration series and embedding in paraffin wax. Cross-sections and longitudinal
sections along the AP and LR axes cut with a microtome were stained with
antibodies against PCNA (Chemicon, Temecula, CA) to visualize proliferating cells
and β-catenin (Sigma, St. Louis, MO) to visualize signaling. Binding of the
secondary antibody and diaminobenzidine (DAB) staining were carried out using the
SK4100 Kit (Vector, Burlingame, CA), according to the manufacturer's protocol as
described (Jiang et al., 1998). Some of these experiments used the Ventana
Discovery (Tucson, AZ) instrument for processing.
2.3.4 In-situ hybridization
In situ hybridization was performed as described previously in Jiang et al. (1998).
Probes for feather keratin A and cytokeratin II were used to visualize differentiating
cells. Feather Keratin A expression was used to assess differentiation status (Yu et
al., 2002). Some of these experiments used the Ventana Discovery instrument for
processing.
26

2.3.5 RT-PCR
RNA expression levels were assessed using semi-quantitative RT-PCR. For this
procedure, the center four tail feathers were microdissected into four components:
the dermal papilla, the ramogenic zone, the collar, and the outer feather sheath. Total
RNA was isolated from selected tissue components using the RNeasy kit (Qiagen)
and cDNA is reverse transcribed using oligo(dT) primers (Invitrogen) and AMV
Reverse Transcriptase (Roche). Primers for hormone biosynthetic enzymes were
described previously (Kamata et al, 2004). All primers used are listed in Table 3.
Control and tested genes were amplified in parallel.

2.3.6 ELISA
DHT levels were analyzed using an Enzyme Linked Immunosorbent Assay (ELISA,
ALPCO) according to the manufacturer’s protocol. The data was read on a
microplate reader (BioTek Instruments ELX-800, available through the USC Dept.
of Pathology) and plotted using Prism3 software.

2.3.7 Hormone and Inhibitor Injections
Local injection of hormone or inhibitor near the tail feathers was injected daily at
1.25 mg/kg/day (DHT), 50mg/kg/day (Flutamide), or the vehicle (1 % ethanol in
corn oil) for control. The effects of DHT and Flutamide on growth rate were tested in
regenerated feathers. Injections were performed for 4 consecutive days followed

27

by plucking. After 5 weeks of continuous daily injections, it is evident that DHT
increased the size of the comb and wattle in both roosters and hens demonstrating
that the hormone is having a systemic effect.

2.3.8 Hormone inhibition by implanting pellets
Another method for hormone and inhibitor delivery was used to save time and
increase the number of birds feasible in the study. Ninety day time release pellets
(Innovative Research) composed of inert control material, Cyproterone Acetate
(CPA-competitive AR inhibitor), Tamoxifen (Tam-competitive ER antagonist), or
Letrozole (Let-an aromatase inhibitor) was implanted subcutaneously behind the
neck of male and female birds. Birds were photographed weekly and blood was
drawn. Upon completion of the experiment, birds were sacrificed and tissues
harvested for further characterization.

2.3.9 Microarray
We compared the expression profiles of male and female collar and dermal papilla
regions from the center 4 tail feathers. RNA was extracted from harvested tissue
using the RNeasy kit (Qiagen) and run on an Affymetrix chicken microarray chip.
The chips were run by our Microarray Core Facility located at Children’s Hospital of
Los Angeles. The data was analyzed with dChip and Array Assist software
(Stratagene). Each array was normalized to all of the chips used in the data set.
Cluster analysis was be used to sort out genes that are differentially expressed
28

between the male and female components. Molecular pathways with multiple
members affected became high priority candidates for further testing.

2.4 Results
2.4.1 Different growth kinetics in male- and female-type tail feathers
Feathers are lost through seasonal molts and regrown throughout a bird's life. This
process can also be initiated by removing the feathers through plucking. After
plucking, a specialized region of the mesenchyme, the DP, is left behind. The
epithelia surrounding the plucked regions represent a wound. The epithelium
becomes organized in the healing process so that a single layer of epithelium covers
the DP by 2 days after plucking. This then forms a stratified epithelium by day 3.
Plucking resets the molecular and cellular processes to a naïve state and
synchronizes ensuing feather growth, enabling direct comparisons between similarly
aged specimens. This model was used to explore the role of hormones in feather
growth.

When the sizes of two epithelial organs A and B are different, it can be due to
differences in growth rate (A>B), or because the duration of the growth period is
longer in A than B (Fig. 3). The difference between male and female tail feather
growth kinetics was determined by plucking the four center feathers showing the
most dramatic sexually dimorphic elongation patterns. There are 14 rectrice tail

29

feathers (Lucas and Stettenheim, 1972). For each bird, the four rectrices closest to
the midline were measured since they have the most pronounced sexual dimorphic
phenotypes. Both male and female tail feathers begin to emerge from the invaginated
feather follicles about 10 days after plucking. Male tail feathers have a much longer
growth period than female tail feathers (males—9 weeks; females—5 weeks), but
they both exhibit similar growth rates during the elongation phase of the feather
cycle (Fig. 4). Male tail feathers have a wider diameter than female tail feathers.
Their calamus, the lower feather portion that remains unbranched, is longer than in
female feathers.
30

Fig. 3 How does the growth of male tail feathers differ from female tail feathers? Do
they (A) grow at different rates or (B) grow at the same rate for different durations?
Black, male; gray, female.
31

Fig. 4 Growth kinetics of male and female tail feathers. Regenerative tail feather
length after tail feathers were plucked (time 0). Note that the male and female
feathers initially grow at similar rates, but the rates diverge at 4-5 weeks. The male
tail feathers then continue to grow for a longer period of time than the female tail
feathers. The increased standard error detected in the male growth curves at later
time points is attributed to a growth gradient due to differences in the growth period
between the center (which grows longest) and lateral tail feathers.
32

2.4.2 Comparison of feather follicles in male and female rectrice feathers
We next examined the feather follicles. Following feather bud development
(Widelitz and Chuong, 1999), the epithelium invaginates into the dermis to form a
follicle structure (Lucas and Stettenheim, 1972; Yu et al., 2004). Thus, the
epithelium forms a cylindrical or tube structure with mesenchyme wrapped inside
(Prum, 1999, Fig. 5). Cells proliferate in the proximal follicle, but start to form barb
branches in the distal follicle that differentiate to become the feather filament, which
is pushed further distally. We have earlier identified a shift of localized growth zones
in these changing morphogenetic processes (Chodankar et al., 2003), and show that
the Wnt pathway (Chang et al., 2004a), along with other pathways, are involved
(Widelitz et al., 2003). Those studies were carried out with flight feathers and body
contour feathers. Here, we examine rectrices (tail feathers) from mature male and
female chickens.
33

Fig. 5 Schematic of feather follicle structure. Feathers initially form as a cylinder.
The mesenchymal structures (shades of red) are the dermal papilla (dp) and the pulp.
Blood vessels (bv) bring nutrients to the growing feather follicles. The epithelia
(shades of blue) will form the feather. The epithelial localized growth zone (LoGZ)
is near the base of the feather above the dermal papilla. Feather branching into barbs
takes place in the differentiating ramogenic zone (rz). After differentiation, the pulp
cells die, enabling the feather branches to open up.

34

The tail feathers of 1-year-old birds were plucked and the feather follicles were
removed at various time points of regeneration. H&E staining was performed on
regenerating adult feathers at various time points. Differences in the size of male and
female 6-week-old feather follicles can be seen (Fig. 6). Data reveal that there are a
few structural differences between male and female tail feathers. Male tail feathers
are not only longer, they are also wider in diameter and contain a larger collar (site of
active proliferation) than female feathers of the same age (Fig. 6).
35

Fig. 6 Molecular expression in male and female tail feathers. Longitudinal male and
female tail feather sections (in regeneration) were stained for H&E, PCNA, Feather
Keratin A (Ker A), Cytokeratin II (Cyto II), and β-catenin (Beta-Cat). The
distributions of PCNA and β-catenin protein were determined using immunostaining.
The distributions of Feather Keratin A and Cytokeratin II were determined using in
situ hybridization. A high-power view of the feather collar region is shown (H&E,
PCNA, and Beta-Cat). Since there was no staining in the collar region for Ker A, the
ramogenic zone is shown for Ker A and Cyto II with size bars=250 μm. A lower-
power view is shown in the upper insets with size bars=0.5 mm and a higher-power
view is shown in the lower insets with size bars=200 μm.
36

37

Regions of proliferating cells were determined by staining for PCNA (Fig. 6). PCNA
staining illustrates the transiently amplified (TA) cells that give rise to the feather
barbs and ridges. Proliferating cells are present at a higher concentration and are
retained for a longer duration in male birds. This proliferation could be the result of
continuous hormonal stimulation in the male birds, resulting in a longer proliferation
period and therefore longer tail feather growth. Feather keratins are not expressed
until keratinocytes reach differentiation zones (ramogenic zone). Feather
keratinocytes in both the collar and feather filament show expression of cytokeratin
II (α keratin; Chodankar et al., 2003; Fig. 6). Based on these staining patterns, we
can estimate the size of the TA cell and differentiated cell compartments in hormone-
sensitive "male"- and "female"-type tail feathers, and hormone-insensitive body or
flight feathers. Furthermore, by observing the formed feather, we see that the males
have a longer calamus, and the dermal ridges begin higher up in the male feather
than the female, which further illustrates structural differences between male and
female birds.

2.4.3 Comparison of feather branches rectrice feathers
As feather filaments extend more distally, they form feather branches through
differential apoptosis (Chang et al., 2004b). The length, angle, and spacing of barbs
can be different in diverse feather types (Prum and Brush, 2002). Aparicio et al.
(2003) report that in some species, male tail feathers sacrifice structural
integrity for enhanced length. We also noted structural changes between the tail
38

feathers from male and female chickens and try to define these parameters as
markers for male or female tail feathers for future studies (Fig. 7).

39

Fig. 7 Morphological difference in male and female tail feathers. H&E staining of
male (A) and female (B) feathers at the same height up from the base reveals
differences in diameter, rachis width, and barb density. The rachides shown here
have regenerated for 9 weeks. The female rachis has finished growing and is larger
than the male rachis at this point. The male rachis will continue growing for an
additional 3 weeks and will ultimately be larger than that of the female. The barb
density at the point of insertion into the rachis is distributed as a gradient in male
rectrices but is constant in female feathers as seen in scanning EM photographs. In
males the density is lowest near the distal tip (compare C, D), moderate in the middle
(compare E, F) and highest at the base (compare G, H) as shown in scanning EM
photographs. Overall, the rachis is wider at the proximal base of tail feathers, but
tapers toward the distal end; however, the width of the male is larger than the width
of the female feathers at equivalent locations. These differences are shown
schematically for male (I) and female (J) tail feathers.

40

41

Our data indicate that male and female feathers in hormone-responsive and hormone-
insensitive regions share similar basic structures, although the size of some of the
features may be increased in hormone-sensitive male feathers. We compared feather
diameter at the widest point, rachis diameter, total feather length, the shape of the
rachis, barb density, location, and number of proliferating cells using image analysis
(Scion Image, Frederick, MD; NIH image software for the PC). Chickens have seven
pairs of tail feathers. Each of the female tail feathers is of approximately the same
length (center feathers 19.09±0.79 cm, most lateral feathers 16.95±0.7 cm), while the
male tail feathers form a growth gradient, with the longest feathers near the midline
(40.49±5.96 cm) and the shortest feathers at the lateral sides (19.65±0.64 cm, n=7)
(Table 1). For this reason, our focus in this study was on the four central tail feathers.

42

Table 1 Tail feather physical characteristics—male versus female
Male tail feather Female tail feather
Length (base to tip) 40.49 ± 5.96 cm 19.09 ± 0.79 cm
Rachis diameter * 350 ± 30 μm 250 ± 40 μm
Barb density index
#
0.68 1
*The difference in male and female tail feather diameter can clearly be seen at the
level of the ramogenic zone.
#
The barb density index = barb density in the distal vane/barb density in the
proximal vane.

2.4.4 Determining the effects of hormone modulation by local injection
Since many types of cancers (breast, prostate, etc.) are hormone dependent it is
necessary to identify the molecular pathway involved in growth after administration
of steroid hormone agonists or antagonists. Perturbation of the estrogen/androgen
ratio by altering a part of the hormone biosynthetic pathway (Fig. 2) should pose
insight into hormone dependent epithelial growth upon the change of morphology in
the feathers. By learning how nature regulates this hormone dependent growth, we
may find pathways that can be exploited to develop inhibitors in order to halt tumor
growth.

43

First, the serum hormone levels in male and female birds were measured. We began
by analyzing the levels of DHT in the blood of the white leghorn birds using an
ELISA for DHT (ALPCO). This in fact determined two things. The first is that the
ELISA can work in birds and the antibody it uses cross reacts with hormones from
this species and the second is that the males do have a higher level of circulating
DHT in the blood similar to humans (Fig. 8)

Fig. 8 DHT ELISA. Serum samples from male and female birds were analyzed using
the ALPCO DHT ELISA kit. As shown, DHT levels in the circulating blood were
significantly higher in the male birds compared to female birds.

44

Hormones and inhibitors were injected locally into the feaher follicles using the
previously described plucked / regenerating feather model (Yu et al., 2002). Topical
administration of the inhibitor onto the surrounding epithelium of the feather follicle
was also done in parallel but showed no significant results. Flutamide (Sigma, CA)
was administered at a concentration of 50mg/kg/day (O’Connor et al., 2002). DHT
(Sigma, CA) was administered by injection at a concentration of 1.25mg/kg/day
(Yoshimura et al., 2003). At various time points feathers were photographed and the
morphology, histology, and molecular expression analyzed (Fig. 9).
45

Fig. 9 Phenotype of male and female birds injected with Flutamide or DHT. Daily
injections of DHT showed the greatest results. As shown, upon injection with DHT
the female birds comb and wattle began to grow. Similar to the females the male
comb and wattle also grew larger and the tail feathers actually began to differentiate
sooner.

46

These initial experiments were performed on juvenile birds. It is entirely possible
that these experiments which were carried out before the animals became sexually
mature, may have resulted in a change in the development of the bird.

We then tested the role of androgen pathway stimulation and suppression with the
experiments described above in adult white leghorns. Local injections of DHT and
Flutamide were administered on a daily basis and the birds were photographed
frequently. Upon visual inspection of the birds, one could first see that the loss of
feathers was apparent. Feathers lined the bottom of the cage on a daily basis
beginning about 2 weeks after the injections began. Photos were taken of the birds
which demonstrate the loss of feathers (Fig. 10). Flutamide injections showed very
minor loss of feathers in both sexes which did not begin until the sixth week of
injections (data not shown). These results suggest that the “hormone non-responsive”
feathers may be able to respond to hormone stimulation.
47

Fig. 10 DHT induced alopecia in birds. Daily injection of DHT caused both male
and female birds to lose feathers in various parts of the body including the neck
region (a), body (b), breast (c, f), saddle (d), and hackle (e).

48

2.4.5 Hormone pathway inhibition studies
We next inhibited hormone action by using implantable time release pellets
(Innovative Research). Ninety day time release pellets for cyproterone acetate
(CPA)-an AR inhibitor (10mg pellets for females and 20mg for males), Tamoxifen
(Tam)-an ER antagonist (25mg pellets for females and 50mg for males), and
Letrozole (Let)-an aromatase inhibitor (10mg pellets for females and 20mg for
males), along with control pellets were implanted subcutaneously behind the neck of
both male and female birds. Placement of the pellets behind the neck prevented the
birds from being able to remove them. The skin behind the neck is also loose, which
enabled us to create just a minor incision that could be closed with a single suture.
There were five male and five female birds per group. We used 2 separate control
groups because the inert ingredients for the CPA pellets were different from that of
the Tam and Let pellets. This gave us a grand total of 50 birds at the beginning of the
experiment. Color coded bands placed around the feet of the chickens enabled us to
keep track of individual birds within the control and treatment groups. A few of the
birds were sick upon arrival from our local farm. I shuffled those to the control
groups and unfortunately 3 males from control group 2 and one female from control
group 1 died during the experiment.

Results from the 3 month inhibition study did not go as predicted. We had hoped to
find a conversion from male to female or vice versa but the results were not so

49

clear cut. Fortunately the Control birds had no phenotypic change as expected. In
fact the female birds in all treatment groups showed no significant phenotypic
changes compared to controls. CPA treated birds, which should have had an
inhibition in AR signaling also showed no significant phenotypic change. I believe
that this is probably due to the fact that the White Leghorn chickens have no
pigmentation present. Based on the literature, injections of estradiol caused a change
in pigmentation in both male and female brown leghorn birds (Juhn 1930) suggesting
that pigmentation is under the control of androgens. In addition to pigmentation, the
size and shape of the feather also resembled that of a female hen after addition of
exogenous estradiol. Taken together this suggests that estradiol is responsible for the
phenotype of the female feather. Molecularly, how is this taking place and what
would happen if we inhibit the conversion of androgen to estrogen by inhibiting the
function of aromatase? Administration of the Let pellets also had no significant
change in phenotype. Aromatase is required to convert androgens into estrogens.
Since Letrazole is an inhibitor of aromatase, I suspect that the Let pellets retained the
male rectrices in their male phenotype. Females already have plenty of estradiol
circulating in their blood streams, so the local inhibition of Aromatase would not
change the phenotype. The Tam treated birds also showed no real change. Since
Tamoxifen is an ER antagonist, I would have expected that it would have had a
significant effect on female rectrices, causing them to assume male default (male)
feather phenotype. Perhaps the concentration was not high enough to totally suppress
hormone-receptor interactions.
50

One change that was noticeable was that at the end of the experiment, the males in
the treatment groups, had tail feathers that were still in anagen phase (Fig. 11).
Typically male tail feathers are done growing at the end of 12 weeks and female tail
feathers halt shortly after 9 weeks. Therefore, the length of the tail feathers is directly
controlled by altering hormone levels but not significantly.
51

Fig. 11 Percentage of feathers remaining in anagen following pellet implantation for
12 weeks. As shown, at 12 weeks all female birds had stopped growing. Male birds
on the other hand had a significant increase in the percentage of feathers remaining
in anagen for all three treatment groups compared to controls.
52

So, this suggested that another mechanism must be present locally to produce
regional specificity within each of the different feather tracts allowing each tract to
exhibit a different phenotype.

Next, we performed microarray experiments on both male and female tail feathers.
For this experiment, we used six male and six female birds. We plucked the tail
feathers and allowed them to regenerate for 3 weeks before surgically removing
them. Only the center four feathers from each bird was used for the experiment,
since these are the feathers that grow the longest in the males, and are therefore
presumed to be the most responsive to hormone stimulus. Next, we microdissected
the feather follicles into four distinct compartments, the mesenchymal compartment
or dermal papilla, and three epithelial compartments that include the collar region
which houses the potential stem cells, the ramogenic region that is developing into
barbs and barbules, and the feather follicle sheath. The individual compartments
from the birds were pooled and homogenized using liquid nitrogen and run through a
shredder column (Qiagen). RNA was then extracted using the RNeasy kit (Qiagen)
and tested for gene expression using Affymetrix microarray analyses at the
Children’s Hospital of Los Angeles core microarray facility . The RNA was
quantitated and tested for integrity. Once the samples had passed, 2ug of RNA was
labeled and run on the Affymetrix Chicken microarray chips. We chose to run only
the dermal papilla and collar regions from both male and female birds. The
microarray results were used to identify potential candidates for us to pursue.
53

Among those on the list included various members of the hormone biosynthetic
pathway (Table 2).
Table 2. List of candidate genes with a 5 fold change generated using D-CHIP
analysis following microarray analysis.

M DP/F
DP Mcol/Fcol
M DP
M
Collar F DP
F
Collar ratio 19/17
ratio
20/18 F/M
WPKCI-8 163.00 19.96 6866.94 6916.83 0.02 346.54 75.34
FZ-8 3.70 3.70 18.78 30.87 0.20 8.34 6.70
WNT2B 98.45 159.56 147.29 19.30 0.67 0.12 0.65
CYP17A1 97.43 33.63 42.80 4.81 2.28 0.14 0.36
COL3A1 7040.64 185.53 7503.53 28.62 0.94 0.15 1.04
SREBF1 36.57 33.51 3.70 58.35 9.88 1.74 0.89
SREBF1 19.99 6.69 3.70 3.70 5.40 0.55 0.28
Netrin G2 27.78 11.69 3.70 21.12 7.50 1.81 0.63
SOX9 20.74 10.46 3.70 20.79 5.60 1.99 0.78
MYB 3.70 21.90 24.28 25.50 0.15 1.16 1.94
FGF-8 3.70 12.27 18.91 26.24 0.20 2.14 2.83
MYO3A 3.70 3.70 19.96 3.70 0.19 1.00 3.20
COL4A4 3.70 50.75 27.02 43.54 0.14 0.86 1.30
COL4A3 3.70 32.95 20.11 9.87 0.18 0.30 0.82
Cadherin 12 3.70 218.37 24.96 203.94 0.15 0.93 1.03
CYP1A1 5.38 18.00 12.33 11.84 0.44 0.66 1.03
Aromatase 7.86 7.30 13.42 15.38 0.59 2.11 1.90
HSD17B4 1379.52 1865.40 1161.34 1061.19 1.19 0.57 0.68
HSD3B1 22.41 62.24 8.50 12.78 2.64 0.21 0.25
HSD3B1 7.98 60.45 15.08 31.00 0.53 0.51 0.67
HSD17B7 236.85 576.40 267.21 635.51 0.89 1.10 1.11
HSD17B11 252.13 466.77 419.48 376.54 0.60 0.81 1.11
HSD17B7 114.85 425.64 119.41 460.77 0.96 1.08 1.07
ESR2 49.06 53.84 36.82 65.65 1.33 1.22 1.00
ESR1 52.31 16.34 25.97 12.71 2.01 0.78 0.56
IGF1R 1069.42 762.47 1199.56 713.09 0.89 0.94 1.04
IGF1 50.12 28.90 61.48 21.98 0.82 0.76 1.06
AR 38.85 36.83 66.87 26.96 0.58 0.73 1.24
SRD5A1 24.36 671.56 21.95 601.97 1.11 0.90 0.90
AHCTF1/SRD5A2 26.39 40.25 3.81 19.20 6.93 0.48 0.35

54

2.4.6 Local conversion of hormones ensures a female phenotype
To verify the distribution of interesting candidate genes that our microarray results
identified, we performed RT-PCR using the primers listed in Table 3. These include
the hormone biosynthetic enzymes Aromatase (Aro), cytochrome P450 side chain
cleavage (P450scc), cytochrome P450 17alpha-hydroxylase/C17-20 lyase (P450c17),
3β-hydroxysteroid dehydrogenase (3B-HSD), and 17β-hydroxysteroid
dehydrogenase (17B-HSD), steroid receptors including the Androgen receptor (AR),
Estrogen receptor type 1 or alpha (ESR1), Estrogen receptor type 2 or beta (ESR2),
and insulin growth factor 1 (IGF1) and its receptor (IGFR1) which are known to act
downstream of the AR in the hair. These genes were compared to beta-actin as a
control. Results are summarized in Figure 12.

Table 3. List of RT-PCR primers.
Gene Sense Antisense Length Reference
AR CCTGAATGAACTTGGGGAGA CCAGACAGGATCTTGGGAAC 618
Aromatase CTCGATTTGGGAGCAAGCTTG GACATTCTCAGCAGTCAGA-
TCTCC
547 Villalpando et
al., 2000.
ESR1 CAAGGCACTGAGCTGGAGA CTGTAGAAGGCTGGAGGAG 380
ESR2 GTGACTGTACAAGCCCAA CATCCAGCAGCTTTCCAA 593
3B-HSD GGCAAGTTCCAGGGCAAGA ACAGGTCACAAGCACGCCT 408 Kamata et al.,
2004.
17B-HSD GGCCATGAGAGCAGTGTTT AGTACACGGCGTTGAAGGG 152 Kamata et al.,
2004.
P450scc TGCTCCCATGCTCTCCAGG GCGGTAGTCACGGTATGCC 382 Kamata et al.,
2004.
P450c17 GCTGCTGAAGAAGGGGAAG CGTCGGTAGGAGGAGTTGA 315 Kamata et al.,
2004.
SRD5A1 GGAGCCAACTTCTTTGGA GGTGACTGTGTCATAGCA 526
IGF1 GATGCACACTGTGTCCTA ATATCAGTGTGGCGCTGA 318
IGFR1 ACACTCCGATGTCTGGTCTT TTGTGTCCTGAGTGCTTGTC 389
Beta-Actin GCCAACAGAGAGAAGATGAC CACAATTTCTCTCTCGGCTG 288 Kamata et al.,
2004.

55

Fig. 12 RT-PCR results for male and female feathers compartments. Of note,
Aromatase is increased the epithelium of the female birds. While many of the genes
are only mildly different between the sexes, SRD5A1 is absent from the dermal
papilla in the females.

The local level of sexual steroids capable of acting on the feather depends upon the
56

presence of androgen and estrogen synthesizing enzymes. Aromatase undoubtedly is
present at a higher level in order to convert any testosterone present in the blood to
estrogen locally in the skin and feathers of the female birds. Additionally, females
also have reduced levels of SRD5A1 especially lacking in the dermal papilla, which
ensures that testosterone is not converted into DHT. ESR1 also appears to be
increased in the derma papilla yet absent from much of the epithelium in the males.
ESR2 appears to be present in different compartments between males and females.
The combination of estrogen receptors and their presence and absence in various
compartments ensures that estrogens act to make the feather more female in normal
female birds. The enzymes 3B-HSD and 17BHSD are also increased in the female
samples compared to males further increasing the amount of estrogens converted
locally in the skin and feather. P450scc and P450c17 did not show that much of a
change. This is probably due to their action being further upstream in the hormone
biosynthetic pathway. IGF1R and IGF1 also did not appear to be significantly
changed albeit IGF1 is expressed at high levels in the dermal papilla of both sexes
compared to the epithelial components. Testosterone and DHT act through the same
nuclear receptor, the AR, with DHT being the more potent ligand. The AR is present
at a higher level in the males than the females, and while it appears relatively absent
from the dermal papilla, its function lies in the epithelial compartment producing
pigmentation. These results are opposed to data known in the hair that suggest the
AR is present in both the dermal papilla and epithelium and signaling occurs through
the dermal papilla to the epithelium. This difference probably occurred as a
57

result of the epithelial appendages of birds having evolved separately from that of
mammals. Another plausible explanation for this is that the epithelial appendages are
performing different functions for the animal. In the birds, the feathers are required
for attraction of mates, warmth, and even flight whereas the hair in mammals
provides warmth and protection.

So we now realize that in these birds the regional specificity is caused by local
conversion of the hormones. Higher levels of estrogens synthesized as well as
converted locally ensure that female birds are phenotypically female since the male
sex is the default sex of birds.

2.5 Discussion
2.5.1 Sex hormones in epithelial–mesenchymal interactions
Humans show regional specificity in their response to sex hormone stimulation, i.e.,
androgens stimulate the growth of beards but also induce male pattern baldness in
the scalp hair. In chicken, feathers also come in different sizes and shapes. Body
feathers (or contour feathers) are small and do not respond well to circulating levels
of sex hormones. Tail feathers grow longer in roosters than hens, suggesting that
their growth is regulated by hormone-dependent pathways (Fig. 2). The mechanism
of their action was not fully understood. We originally devised three possibilities
describing possible ways that sexual dimorphic feathers can occur (Fig. 13). In
58

model "a", the default tail feather prototype in the absence of hormone stimulation is
a body feather. Estrogen-mediated events transform the prototype to the "female-
type" tail feather; androgen-mediated activities transform it to the "male-type" tail
feather. In model "b", the default tail feather form is the "female" tail feather, which
is transformed by androgen stimulation to the "male type". In model "c", the default
tail feather type is the "male" type, in which estrogen transforms to the "female
type". From past literature and our own data we can conclude that model "c" is the
correct model. The male sex is the default and thus the male tail feather is also the
default. Upon addition or local conversion of estrogen, estrogen signaling occurs
through the dermal papilla and causes the differentiation and cessation of
pigmentation which in turn leads to the formation of a female feather.
59

Fig. 13 Possible models of hormone effects. Model A, the body (contour) feather is
the default type. Sex hormones influence whether a "male" or "female" feather will
develop. Model B, the "female" is the default feather type. Androgens cause a
transition to a "male" feather type. Model C, the "male" feather is the default and
estrogen converts the feather to a "female" feather type.

60

3B-HSD converts DHEA into androstenedione. In humans, two isoforms have been
described, but the skin seems to only express the type 1 isoform (Zouboulis 2007).
3B-HSD is mainly located in the sebaceous glands of humans whereas in the
chicken, since they lack sebaceous glands, 3B-HSD is seen weakly in the dermal
papilla and more strongly in the epithelial components. Androstenedione is further
converted to testosterone by the enzyme 17B-HSD. In hair follicles, 17B-HSD is
localized in outer root sheath cells (Zouboulis 2007). In the chicken, 17B-HSD is
also located in the epithelial components of the feathers.

Estrogens exert their effects by binding to two receptors that have been identified.
ERα and ERβ are encoded by two different genes. They differ in size; ERα is 67 kDa
and ERβ is approximately 54 kDa (Rollerova and Urbancikova, 2000). The gene for
ERα is localized on chromosome 6 and that encoding ERβ is found on chromosome
14 in humans (Enmark et al, 1997). ERα has a higher affinity than ERβ for17 β-
estradiol, the most active estrogen form. In human skin, ERα and ERβ have different
distributions. ERα was only found in the sebaceous gland, while ERβ was found in
the outer root sheath, epithelial matrix, and dermal papilla (Thornton et al, 2003).

The principal action of androgen is to regulate gene expression through the AR,
which belongs to the superfamily of nuclear receptors. Androgens (testosterone and
DHT) control the development, differentiation, and function of male reproductive
and accessory sex tissues, such as the seminal vesicle, epididymis, and prostate.
61

Other organs and tissues, such as the skin, skeletal muscle, bone marrow, hair
follicles, and brain, are also under the influence of androgens. Androgens are
primarily synthesized in the gonads of chickens but can also be made in the
epididymis following partial or complete castration (Budras and sauer, 1975). The
AR has also been mapped to a number of chicken tissues, including the skin
(Shanbhag and Sharp, 1996). In human skin, the AR was found in the dermal papilla
of the hair follicle (Randall et al, 1992) and the basal cells of the sebaceous gland
(Thornton et al, 2003; Pelletier 2004). Androgen receptors are found in higher levels
in balding dermal papilla cells of the hair follicle than those in the non-balding scalp
(Hibberts et al, 1998). The growth of sex hormone responsive hairs is dependent
upon the binding of testosterone or DHT to the AR. In humans, 5α-Reductase
expression is predominantly in the sebaceous and sweat glands, although activity is
seen in epidermal cells and hair follicles but to a lesser extent (Zouboulis 2007).

In humans, androgens appear to act mainly through type 2 5α-reductase and the AR
on dermal papilla cells in the hair (Zouboulis 2007). Whereas in chickens, since the
AR is located in the epithelium and not the dermal papilla, the presence of 5α-
reductase is likely present in all tissues order to convert testosterone into DHT to
activate pigment production in the feather. The exception to this is that 5α-reductase
62

is absent from the female dermal papilla to further ensure that higher levels of
estrogen are present to produce a female feather.
2.5.2 Potential cellular/molecular mechanisms
It has long been known that male tail feathers grow longer than female tail feathers
but how they do this is unknown, particularly at the cellular/molecular level.
Presumably, female chicken feathering patterns result from ovarian estrogens and
male feathering patterns result from androgens. In our recent work, we used the
concept of a localized growth zone to explain the shape of feather/scale (Chuong et
al., 2000), chicken/duck beaks (Wu et al., 2004), and liver (Suksaweang et al., 2004),
etc. successfully. Mainly, the duration, size, and number of localized growth zones
can shape a growing organ. While it is not clear which hormones are causing the
effects, here we show that the sex steroids do have a different impact on the levels of
activity and duration of localized growth zones in male and female tail feathers (Fig.
14). These differences also occur elsewhere within the integument, including the
comb, wattle, spurs, hackle feathers, and saddle feathers. In each case, these features
are larger in males than females. Sexual dimorphisms appear developmentally early
in the comb, wattle, and tail feathers and later in the spurs, hackle feathers and saddle
feathers (Jacob and Mather, 2000). Since the tail feathers and other secondary
characteristics grow more robust in the male, we presume that these differences are
due to hormonal control. Results of the growth kinetics study indicate that male tail
feathers grow at a similar rate but possess a considerably longer growth period

63

than those of the female. This change in duration must mean that particular
molecules controlling this growth phase are influenced by sex hormones.

Fig. 14 Sex hormones' influence on feather diameter and length. The dermal papilla
(red), LoGZ (light blue), and differentiation zone (DZ, or ramogenic zone, dark blue)
are shown.

64

One thought for the difference in growth is that the males sacrifice structural
integrity for length. This would mean that there would be differences in the feather
morphology of the feathers, which we noted. By careful observation of the tail
feathers, we noted that male tail feathers are longer, have a larger degree of
curvature, have a wider feather diameter, a longer calamus, and the dermal ridges
begin higher up in the feather than in the female. These structural differences may
reflect differences in signaling pathways. We tested molecules known to be involved
in the growth pathways of other tissues as well as molecules that would identify
possible differences in proliferation and differentiation. Results indicate that
proliferating cells are present at a higher level and are retained for a longer duration
in male birds. This proliferation could be the result of continuous hormonal
stimulation in the male birds, resulting in a longer proliferation period and therefore
longer tail feather growth.

Sexual dimorphism of feathers may result from modulating the shared stem and TA
cell configurations and underlying molecular pathways. TA cells might have the
ability to mediate differences in proliferation causing the sexual dimorphism of
chicken tail feathers. Since males and females have different levels of hormones
present throughout development, a comparison of the size and distribution of TA cell
populations and dermal papilla cells will help to identify which cell groups may be
the target of sex hormones. These observed paradoxical effects of sex hormones on
feather growth suggest that there are complex cross talk mechanisms and
65

highlight the importance of having an in vivo model so that no components are
missed, as well as a normal model so we that know how they are regulated normally.

How do hormones exert their effects on growth? We believe that they modulate
growth regulatory pathways that are also present in hormone-insensitive tissues. A
number of growth pathways involved in feather morphogenesis have been described.
Our previous work has focused on the role of the canonical Wnt–β-catenin signaling
pathway in feather growth. Wnts, which invoke the canonical β-catenin pathway,
induce new feather growth from scales and apteric regions (Noramly et al., 1999;
Widelitz et al., 2000). In feather-producing regions, these Wnts induce enlarged and
distorted feather phenotypes (Chodankar et al., 2003). Blocking Wnt signaling with
Dkk suppressed feather growth (Chang et al., 2004). Additionally, β-catenin can bind
directly to the androgen receptor to enhance transcription synergistically (Truica et
al., 2000). We recently found that β-catenin can synergistically act as a co-activator
or the androgen receptor with co-activator-associated arginine methyltransferase
(CARM) 1 (Koh et al., 2002). Therefore, there is good reason that sex hormones may
modulate growth via the β-catenin pathway. Another candidate is the bone
morphogenetic pathway (BMP) pathway. BMP7 has been shown to be highly
regulated with estrogen receptor levels, which is an important prognostic marker for
breast cancer (Schwalbe et al., 2003). BMPs and their interaction with type II BMP
receptors have also been shown to contribute to the proliferation of human breast
cancer cells (Pouliot et al., 2003). A direct interaction between Smads and ER
66

was shown to cause an inhibitory action on Smad activity, which suggests cross-talk
between BMP and steroid hormone pathways, particularly the ER pathway
(Yamamoto et al., 2002). We are currently examining the role of β-catenin in
mediating hormone-dependent growth to determine how hormones exert there effects
on normal growth-mediating processes.

2.5.3 Regional specificity of sex hormone effects
Why do hormones exert different effects in different regions of the integument? Why
do androgens induce facial hair growth (beard) but suppress scalp hair growth? In
our model system, the question can be rephrased as why do the tail feathers respond
differently to hormone action than the comb and wattle and why are the body
feathers seemingly unaffected? These differential responses are due to regional
specificity. How is this regional-specific hormone response established? Different
molecular machinery in cells within each region may cause different responses to the
same hormonal stimuli. While steroid receptor co-activators and co-suppressors have
made much progress in mammals (Chang, 2002), these studies are still at a primitive
stage in birds. The skin Hox code may be involved in setting up such different
competence in skin regional specificity (Chuong et al., 1990).

Many aspects of hormone metabolism take place in the peripheral tissues. Hence the
local concentrations of hormone forms are dependent on the levels of
circulating
67

precursors within the blood and the availability of local metabolizing enzymes. A
schematic of hormone biosynthesis indicating enzymes (green) and products (black)
is shown in Fig. 2. 3β-hydroxy-steroid dehydrogenase converts DHEA to
androstenedione. It is also involved in converting androstenediol to testosterone.
17β-hydroxy-steroid dehydrogenase converts DHEA to androstenediol,
androstenedione to testosterone, and estrone to estradiol. Aromatase converts
androstenedione to estrone and testosterone to estradiol.

The sex hormone biosynthetic pathways are highly conserved across species. In each
case, the steroids must bind to their receptors, which interact with co-activators and
co-suppressors to modulate transcriptional activation. Many of the molecular aspects
of transcriptional activation have been elegantly demonstrated using tissue culture
cells (reviewed in Chang, 2002). It is thought that similar pathways are active in
mediating similar events in tissues and modulate epithelial–mesenchymal
interactions. Since these processes are so important to diseases of skin appendages, it
is critical to have a model in which to assess hormone effects in a physiological
situation in vivo. We have observed that the chicken tail feathers respond to hormone
stimulation by altering its growth and molecular expression patterns. This
experimental model can now be exploited to obtain new understandings that may in
the future assist patients suffering from hormone-dependent growth-related diseases
affecting other skin appendages, such as hair growth-related diseases, breast
68

cancer, or prostate cancer.
2.6 Summary
Hair, mammary glands, feathers, etc. are all skin appendages derived from the
epidermis as a result of epithelial–mesenchymal interactions. Pelage and skin can be
markedly dimorphic in many mammals including deer and primates. The growth of
these epithelial organs is regulated by fundamental morphogens including BMP,
SHH, Wnt/β-catenin, etc. (reviewed in Chuong 1998; Widelitz et al, 2003). Mutation
or de-regulation of these morphogens has been identified in many human diseases
including tumors and genetic diseases. Sex hormones also play a major role in
regulating the growth and phenotypes of these epithelial organs differently in male
and female animals (sexual dimorphism). How the sex hormones are coupled to the
morphogens is mostly unknown; hence, β-catenin, BMPs, etc., are candidates that
may couple the two pathways. It has been suggested that female chickens have
feathering patterns resulting from ovarian estrogens (George and Wilson 1980),
which in combination with our data and data reported in the literature leads us to
believe that the male feathering pattern is the default pattern and estrogens are the
key to the female feathering pattern. Another way of analyzing the role of hormones
in skin appendage growth is to assess them in various diseases. Examples include
androgenic alopecia, 5α-reductase deficiency, henny feathering, hyperactive mutated
aromatase, etc. The feather offers a unique opportunity to carry out experiments
involving sex hormones and study their effects in epithelial–mesenchymal
interactions, in vivo.
69

Our data, in addition to the classical papers involving caponization and
ovarectomies, taken together suggest that in fact the head furnishings and
pigmentation in the birds are clearly responsive to Androgens. Removal of the ovary
leads to a larger comb and wattle, while removal of the testes leads to the reverse
phenotype, a smaller comb and wattle. Daily injections of estrogen into a male bird
causes pigment production to halt, but can be reversed by removing the source of
estrogen. The plumage is now thought of as being responsive to estrogens. If you
remove the ovary the feathers can now grow long like a male, but if you remove the
testis you end up with either no change in length or in some cases they grow even
longer (Domm 1927).

70

Now coming full circle, it seems that nature has put in place a system in the chicken
that while based on genetics, had led to the male being the default sex in the birds.
Nature has gone out of the way to molecularly ensure that female birds stay female.
This is done by increasing the level of estrogens seen locally in the skin by
modification of the level of hormone biosynthetic enzyme expression in various
compartments of the feather.

71

3.0 Chapter 3 – Mouse Mammary gland project
3.1 Introduction
Mammary glands, like other ectodermal organs, are produced through a series of
epithelial-mesenchymal interactions (Cunha 1992). The mammary gland develops in
three stages including embryogenesis, puberty, and pregnancy (Daniel 1987;
Sakakura 1987; Hennighausen 1998). Mice develop five pairs of mammary glands
which initiate around E10-11. By the time of birth, mammary epithelium branches
into the stromal fat pad. After birth mammary glands undergo three stages of growth:
1) estrogen dependent ductal growth during the 6-8
th
week after birth, 2) pregnancy
stimulated development of lobular-alveolar structures and, 3) remodeling associated
with the loss of alveolar structures after weaning (Strange et al, 1992). Several
molecular pathways, such as EGF, Wnt, FGF and TGF-beta were previously shown
to be involved in the development of mammary glands. Here we explore the role of
the BMP pathway. For this purpose we utilized a transgenic mouse (KRT14-Noggin)
that over expresses Noggin, an inhibitor of the BMP pathway, under the regulation of
the keratin 14 promoter which directs expression to the epithelium including the
mammary gland myoepithelium in which to study the effects of the BMP pathway on
mammary gland morphogenesis.

In the mouse, mammary glands are first observed as five symmetric (left–right) pairs
of placodes visible by around embryonic day 11.5 (E11.5) (Mailleux et al,
2002).
72

Around E14, the epidermis of the male mice, upon stimulation by androgen,
becomes separated from the underlying mesenchyme that is undergoing apoptosis
(Wysolmerski et al, 1998). Contrary, at E15.5 the epithelial bud in female mice
forms a mammary sprout. This mammary sprout then invaginates through the
underlying mesenchyme and at E16.5, a rudimentary ductal tree is already forming.
Subsequently, the nipple sheath begins to form in the overlying epithelium (Foley et
al, 2001). The nipple sheath is then capable of undergoing a thickening process and
subsequent invagination into the underlying dermis. This process is dependent on
PTHrP/PTH-R1 signaling (Foley et al, 2001). The nipple and underlying mammary
gland is one of the sexually dimorphic characteristics present in only female mice.
Proper formation of the nipple and mammary gland are essential for nursing and thus
survival of the pups.

Bone morphogenetic proteins (BMPs) are involved in the development of various
organs including the bone and mammary gland. BMPs were also suggested to control
early mammary gland development (Phippard et al, 1996). SMADs, downstream
mediators of BMP receptor binding, are involved in the epithelial mesenchymal
interaction during development. SMADs have also been implicated in mediating the
epithelial to mesenchymal transdifferentiation in epithelial cells of the breast (Piek
1999). Overexpression of a dominant negative BMP-2 receptor interferes with
BMP-2 induced SMAD 1 transcriptional activity (Pouliot 2003). SMAD 4 is
considered to be a co-mediator SMAD that associates with various receptor
73

SMADs to regulate gene transcription (Piek 1999). Furthermore, SMAD 4, a
downstream member of the BMP pathway, is suggested to interact with the estrogen
receptor (Yamamoto 2002) and may work as a co-repressor (Wu 2003). This cross
regulation between estrogen and the BMP signaling pathways involving SMADs
may have implications in both hyperplasia and tumorigenesis.

In this study, we examined the role of the BMP pathway during the growth and
remodeling of the mouse mammary gland. We demonstrate that BMP signaling is
central to maintaining an organized stroma surrounding the ducts. The stroma is key
to the proper involution of the mammary gland following pregnancy and subsequent
weaning of the pups.
3.2 Methods and Materials
3.2.1 Production/Genotyping of the transgenic KRT14-Noggin mouse
We have been using a KRT14-Noggin mouse model to study the role of BMPs in
mammary gland development. This transgenic mouse model was generated in the
Norris Cancer Center transgenic mouse facility at the University of Southern
California and described in detail in Plikus et al. 2004. Briefly, these transgenic mice
were created by subcloning chicken Noggin into the human keratin 14 promoter
vector. The human KRT14 promoter-chicken Noggin-poly A inserts were released
from plasmid, purified, and microinjected into the male pronucleus of a
74

fertilized egg of C57BL/6J x CBA/J mice. The injected eggs were them re-implanted
into a pseudopregnant C57BL/6J x CBA/J females. Founder KRT14-Noggin mice
were then backcrossed for six generations onto the C57BL/6J background.

KRT14-Noggin

mice were confirmed by polymerase chain reaction (PCR)
genotyping using chicken Noggin-specific

primers: 5'-
CCAGATCTATGGATCATTCCCAGTGC-3' and 5'-
GGAGATCTCTAGCAGGAGCACTTGCA-3' to screen for the presence of the
chicken Noggin transgene. Tail genomic DNA was extracted as described in
manufacturer’s

protocol (Qiagen, Valencia, CA). PCR products were amplified

in
separate reactions using the three-stage PCR program: 94°C

for 3 minutes; 94°C for
1 minute, 55°C for 1 minute,

72°C for 1 minute (30 cycles); 72°C for 10 minutes.

Southern Blot analysis was used to confirm the identities of the founder mice (Plikus
et al, 2004).
3.2.2 Histological, histochemical, and Immunohistological Staining
3.2.2.1 H&E
Specimens were surgically excised followed by fixation in 4% paraformaldehyde
(PFA) buffered with phosphate buffered saline (PBS) overnight at 4
o
C. The samples
were then subjected to a series of dehydration and embedding in paraffin wax.
Sections of the fourth inguinal mammary gland were cut longitudinally with a
microtome at 5um. Standard hematoxylin and eosin (H&E) staining was
75

performed for basic histological analysis.
3.2.2.2 Carmine Aluminum Staining
Whole mount mammary gland analysis was performed according to the protocol
provided by the Laboratory of Genetics and Physiology (National Institutes of
Health). To perform whole mount analysis, the left inguinal mammary glands were
used. Excised tissue was fixed in Carnoy’s fixative for 4 h, rehydrated, and then
stained overnight in Carmine Aluminum solution. The samples were then dehydrated
in an ethanol series and cleared in xylene before mounting and photographing.
3.2.2.3 Immunostaining Sections
Specimens were fixed in 4% PFA/PBS overnight at 4
o
C followed by dehydration
and embedding in paraffin wax. Sections of the fourth inguinal mammary gland
were stained with antibodies against KRT14 (Lab Vision), smooth muscle actin
(SMA) (Lab Vision), and β-catenin (Sigma) to visualize signaling. Binding of the
secondary antibody and diaminobenzidine (DAB) staining were done using the
SK4100 Kit (Vector), according to the manufacturer’s protocol as described
previously (Jiang et al, 1998). Masson’s Trichrome Staining (Sigma) was also
performed using a standard protocol.
3.2.3 In-situ hybridization
In-situ hybridization was performed as described previously (Jiang et al, 1998).
Probes for mouse BMP4 and chicken Noggin were used to visualize BMP
pathway signaling within the ducts of the mammary glands. To detect RNA
76

expression, the tissue sections were hybridized with digoxigenin labeled probes and
the signals
detected using an anti-digoxigenin antibody coupled to alkaline phosphatase. Some
of these experiments use the Ventana Discovery instrument for processing (Ventana
Medical Systems).

3.2.4 Induction of involution
Wild-type and transgenic KRT14-Noggin lactating C57Bl/6 mice had their litters
adjusted to six in number at

parturition because the transgenic mice averaged this
number of pups per litter. Glands were harvested from 8-week-virgin, 2-day-
lactation, and after full lactation was established (10 days of nursing), pups were
removed to initiate involution. The glands were then harvested at 4, 10, 14 days after
forced weaning. Gland morphology during involution was analyzed following H&E
staining.
3.3 Results
3.3.1 Mammary gland development in KRT14-Noggin mice
Typically the mouse mammary gland undergoes a regular pattern of morphogenesis
and differentiation during postnatal development into puberty and pregnancy.
Previous studies indicate that the BMP pathway may be involved in the development
of the mammary gland (Phippard, et al, 1996) and this process may be affected by
interactions with PTHrP (Hens et al, 2007). Presently, no studies have examined
77

the role of the BMP pathway on branching morphogenesis and collagen deposition.
Therefore, the morphology of KRT14-Noggin and wild type fourth inguinal
mammary glands were assessed. Whole-mount Carmine staining of 8-week-old
virgin mouse mammary glands showed that, the overall organization of the ductal
tree was normal in the presence of exogenous Noggin (data not shown). Hematoxylin
and eosin (H&E) stained sections from 8-week-old virgin KRT14-Noggin mice also
showed no significant differences in either the ductal structures or their density (data
not shown). On the other hand, when you compare the ductal structures during the
process of involution, significant changes are observed. The amount of epithelium
fails to apoptose and remains present to a large extent in the KRT14-Noggin mice
compared to controls (Fig. 15). The ducts in the KRT14-Noggin mice also appear to
be dilated compared to controls (Fig. 15).

However, the litter sizes from KRT14-Noggin mice matings were reduced compared
to wildtype, but all pups were normal in size. Greater than 50% of all pups born to
KRT14-Noggin mice died if not placed with a feeding surrogate mother. All of these
findings indicate that KRT14-Noggin mice have undergone normal mammary gland
development but perhaps some defects lie in either the production of milk or the
gland responsible for providing the milk. Another possibility is that the behavioral
instincts of the Noggin mice have been altered but this point will not be addressed
further.
78

Fig. 15 Whole mount Carmine staining of mammary glands. Of note, the large ducts
remain dilated even after 14 days of Involution.

79

Fig. 16 Stromal increase and milk products sustained in the ducts. Evident at 20X
magnification, there appears to be stroma surrounding the duct in the KRT14-Noggin
mouse mammary gland as well as retained milk products within the duct.
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Normal signaling is observed in the mammary glands in KRT14-Noggin mice
Next, we set forth to uncover possible molecular changes within the mammary
glands of the KRT14-Noggin mice in comparison to their wild type littermates at
both virgin and involution stages (Fig. 17). In addition to H&E staining we stained
for Keratin 14 (KRT14), smooth muscle actin (SMA), and beta-catenin. Both KRT14
and SMA stained the myoepithelial cells in both wild type and KRT14-Noggin
mammary glands at both pubertal and involution stages which is the location for
Noggin expression in our KRT14-Noggin transgenic mice. Beta-catenin was positive
in the epithelial cells at all stages. We will continue to screen for differences between
the mammary glands of the KRT14-Noggin and wild type mice.

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Fig. 17 Immunostaining of wildtype and KRT14-Noggin mice. Immunostainingwas
performed on virgin and 14 day involution staged mice for keratin 14 (K14), smooth
muscle actin (SMA) and beta-catenin. All molecules stained in a similar pattern and
there were no significant differences in staining between the mice.
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3.3.2 BMP4 is increased in the mesenchyme of KRT14-Noggin involuting
mammary glands
Since we knew that our KRT14 promoter was driving the expression of Noggin, we
decided to perform in-situ hybridization using probes against Noggin and BMP4
(Fig. 18) As expected Noggin was present in the epithelium of the wild type mice but
over expressed in the KRT14-Noggin transgenic mice at all stages examined (Fig.
18). BMP4 however, was decreased in the virgin stage in the KRT14-Noggin mice
compared to wild type but at the involution stage it re-emerges in the expanded
stromal tissue and remains negative in the ductal epithelium. The reason for this
phenomenon has yet to be uncovered but suggests the presence of a regulatory
feedback loop.
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Fig. 18 In-situ hybridization of the mammary glands of wild-type and KRT14-
Noggin +/+ mice. Noggin is overexpressed in the epithelium of the KRT14-Noggin
mice as expected. BMP4 was decreased in the epithelium of the virgin Noggin
overexpressing mice but during involution begins to be expressed by the stroma
compared to wild-type controls.
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3.3.3 Delayed mammary gland involution in KRT14-Noggin mice
Mammary glands undergo the process of involution when the suckling stimulus from
their young is lost. Most of the involution process occurs during the first four days
and typically lasts on average 10 days in mice (Lund 1996; Monks 2002). The
process of involution has been reported to occur in two stages, the first which is
reversible, lasts for 48 hours after weaning in which the mammary epithelial cells
lose their differentiated function. During the second phase, the extracellular matrix,
including the basement membranes in the mammary glands are degraded by
proteases, leading to the loss of the lobular-alveolar structures. As the process of
involution continues, mammary epithelial cells disappear and adipocytes
concomitantly reappear to occupy the mammary gland.

To identify the physiological roles of the BMP pathway on mammary gland
morphogenesis and remodeling we chose to macroscopically and microscopically
characterize the mammary gland of KRT14-Noggin mice through pregnancy,
lactation, and at three different time points during involution. We anticipated the
data from puberty would help us to understand the roles of the BMP pathway in the
developmental process of the mammary gland and data from pregnancy and
involution would give us insight into the roles of the BMP pathway during
remodeling. To explore the effects of the loss of BMP pathway activity during
mammary gland involution, pups were removed from their mothers at day 10 of

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lactation, and the mammary tissue was collected from 8-week-virgin, 2 day lactation
(0-day-involution), 4-day-involution, 10-day-involution, and 14-day-involution. Fig
19 shows H&E stained sections of wildtype and KRT14-Noggin mammary glands
during lactation and involution. At day 2 of lactation (0-day involution), the majority
of the mammary gland is composed of alveoli lined by epithelial cells that can
secrete milk and there was no phenotypic difference between wildtype and KRT14-
Noggin mice. As involution progressed, the wild type mice exhibited a cellular
pattern in which most of the lobules and alveoli had collapsed by day 4 of
Involution.

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Fig. 19 H&E staining of wild-type and KRT14-Noggin involution staged mammary
glands. Of note are the enlarged ducts present during involution in the Noggin
overexpressing mice. Milk products also appear present.
87

Thus by day 10 of involution, the wild-type mammary gland was remodeled with
only occasional epithelial cords and ducts remaining surrounded by stroma and
adipocytes. In contrast to wildtype mice, the KRT14-Noggin mice demonstrated a
markedly retarded involution process. For example, only a few alveoli in the KRT14-
Noggin mice had began to collapse at day 4 while even at day 10 and 14 of
involution, some alveoli remained intact (Fig. 19) and the gland resembled that of a
day-4 wildtype mammary gland (Fig. 19). Both day 10 and day 14 involution glands
from the KRT14-Noggin glands contain dilated ducts with the presence of milk
lipids. Thus the extent of remodeling of the lobules and alveoli in KRT14-Noggin
mice was considerably slower than in the wild type mice.

3.3.4 Collagen I is maintained in the mammary glands of KRT14-Noggin mice
after involution
We noticed increased extracellular matrix in KRT14-Noggin mammary glands and
sought to identify its components. Trichrome staining was performed following
standard protocols on both wild type and KRT14-Noggin mammary glands and
reveals that the stromal increase visible after H&E staining, consists of collagens
(Fig. 20) and is increased following involution. This collagen deposition is evident
even in the mammary glands of KRT14-Noggin at the 8-week-old virgin stage.
Immunostaining for collagens I, II, IV, V indicate that the majority of the collagens
synthesized are that of collagen I (Fig. 21).
88

Fig. 20 Trichrome staining of wild-type and KRT14-Noggin mammary glands
undergoing involution.
89

Fig. 21 Collagen I staining of wild-type and KRT14-Noggin mammary glands
undergoing involution. Collagen I is the main collagen present surrounding the ducts
in the KRT14-Noggin glands.
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3.4 Discussion
Previous studies indicated that the BMP pathway may normally play a role in mouse
mammary gland development (Phippard et al, 1996) as well as other developmental
processes. The results from our experiments highlight the importance of the BMP
pathway in the remodeling and morphogenesis of the mammary gland and suggest a
possible mechanism by which BMPs affect the involution process and ECM
accumulation in KRT14-Noggin mouse mammary glands. During involution
following pregnancy, the mammary glands of these mice have tremendously
increased amounts of ductal branching. This may be due to increased branching or
decreased apoptosis.
3.4.1 The loss of the BMP pathway via Noggin overexpression delays
mammary gland involution while increasing the amount of collagen I
present
We have also observed an extensive amount of collagen surrounding each duct in the
mammary glands of the KRT14-Noggin mouse. The data presented here suggests that
BMPs may control the amount of collagen deposited within the breast. Collagen
deposition is present in the KRT14-noggin mice beginning at the pubertal stage of the
mouse as shown by Trichrome staining and immunohistochemistry for collagen I. It
has yet to be determined at what developmental stage this collagen is being
deposited.

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Ectopic noggin expression suppresses BMP activity in the epithelium and immediate
surrounding stroma, but BMP expression is increased perhaps in a compensatory
fashion in regions of the stroma further out, away from the myoepithelium.

3.4.2 BMP and Human Disease (Cancer)
In the E13.5 mammary gland, BMP2 is localized in the epithelium and BMP4 in the
mesenchyme (Phippard et al, 1996). BMP-2 levels are decreased in various forms of
invasive, non-invasive, and liver metastatic breast tumors compared to normal breast
tissue (Reinholz 2002). Recently BMP2 has been shown to inhibit MCF-7
proliferation

while inducing p21 expression (Ghosh-Choudhury 2000). BMP2 also
increases the level of PTEN, a known tumor suppressor, in MCF7 breast cancer cell
line (Waite 2003). BMP-2 causes hypoposphorylation of pRb thus inhibiting
proliferation in breast cancer cell lines (Ghosh-Choudhury 2000). BMP-2 is also
found to be expressed in both breast cancer cell lines and primary breast tumor tissue
samples (Clement 2000), The expression of BMP-7 was highly correlated with the
levels of estrogen and progesterone receptors which are important markers for breast
cancer prognosis and therapy (Schwalbe 2003). These new results show that we need
to look more into the roles of BMP in the normal and abnormal growth of mammary
glands. Here we explored how BMPs are involved in mammary gland development,
through puberty, pregnancy and parturition.

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Increased breast density is the single highest risk factor for breast cancer, and yet its
cause remains unknown (Pike 2004). Increased density, often referred to as
mammographic density, has been proposed to consist of cellular and/or extracellular
components. Since the mammary gland is remodeled throughout adult life, the
interaction between the epithelium and extracellular components are constantly in
play. Further verification of the action of these pathways in vivo might suggest a
mechanism by which stromal density is created in human breasts and by identifying
molecules that connect the BMP pathway and tumorigenesis we will understand
more about the possible mechanism of breast cancer development.

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4.0 Chapter 4 – Mouse nipple
4.1 Introduction
The development of diverse ectodermal organs such as mammary glands, feathers,
and hair have much in common and share developmental stages such as induction,
morphogenesis, and differentiation (Chuong 1998). Many ectodermal organs have
even become defining characteristics of specific vertebrate classes; i.e.feathers and
hair are cardinal characteristics of the Aves and Mammalia classes, respectively (Wu
et al, 2004). Although ectodermal organs arise from different developmental
mechanisms, several aspects of their development and morphogenesis run in parallel.
Early stages of development include the formation of a dense mesenchyme
underlying an epithelial bud followed by invagination or evagination coupled to
differential growth. Then the epithelial appendage undergoes a process of
differentiation to unveil the unique ectodermal organ structure and function. Each of
these ecotdermal organs expand the skin surface area in different ways to provide
methods for display, camouflage, feeding of the young, warmth or cooling, flight,
etc.

For this paper we analyzed ectodermal organs which lie along the mammary line. In
the mouse, the differentiation of stem cells into a mammary gland is first observed
by the appearance of five symmetric (left–right) pairs of placodes visible by
scanning electron microscopy at embryonic day 11.5 (E11.5). These are seen as

94

elevations above the surrounding surface ectoderm (Mailleux 2002). At this stage the
epidermis of male mice is under the influence of androgens and the epithelial bud
becomes separated from the epidermis as a result of apoptosis in the underlying
mammary mesenchyme around E14 (Wysolmerski et al, 1998). Whereas in their
female counterparts, at E15.5 the epithelial bud forms a mammary sprout and
invaginates through the underlying mesenchyme. This is followed at E16.5 by the
formation of a rudimentary ductal tree derived from the branching of the epithelial
sprout into the mammary fat pad and the induction of the nipple sheath in the
overlying epithelium (Foley et al, 2001). Thus male mice fail to form both mammary
glands and nipples. The nipple sheath results from epidermal thickening which will
invaginate down into the underlying dermis forming a halo surrounding the location
of the mammary sprout and is formed as a result of PTHrP/PTH-R1 signaling (Foley
et al, 2001). The nipple is considered a type of specialized epidermis that shows
distinct patterns of differentiation and keratin expression in order to withstand the
mechanical strain of nursing the pups (Eastwood, 2007).

In normal mice, hair follicles do not develop in nipple forming skin but they are
found in numerous regions covering the body surface. In these regions proliferating
keratinocytes can systematically differentiate giving rise to the basic structure of the
hair follicle and this leads to the formation of various structures in the hair follicle
including the inner (IRS) and outer root sheath (ORS), medulla, cortex, and central
cuticle of the hair shaft. The epithelium can further differentiate into a
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sebaceous gland which is comprised of lipid filled sebocytes that release their
contents (sebum) onto the surface of the epithelium. Stem cells have been proposed
to lie in a specialized region of the hair follicle known as the bulge and are said to be
responsible for replenishing the sebaceous gland in addition to generating the hair
lineages (Taylor 2000;Oshima 2001).

Multiple signaling pathways have been implicated in various stages of the hair
follicle cycle including Wnt, fibroblast growth factor (FGF), sonic hedgehog (Shh),
and the TGF-beta superfamily signaling pathways. The BMP pathway consists of
more than 20 secreted growth factors and their receptors that are involved in many
aspects of development. Various BMPs and secreted antagonists of the BMPs such
as Noggin, Chordin, and Follistatin are of particular importance for hair follicle
morphogenesis and cycling. BMPs have been described as being classical
morphogens because they have the ability to alter cell fate (Gurdon, 2001; O'Connor
2006; Rosen 2006). BMPs such as BMP2, 4, and 7 signal through a heteromeric
complex of type I and II receptor serine/threonine kinases (Massague 1996). Binding
of the BMPs to the receptors induces the phosphorylation of the receptor regulated
SMAD family (rSMADs) members (SMAD1,5,8) that then associate with co-SMAD
(SMAD4). This phosphorylated complex can then translocate into the nucleus and
regulate the transcription of various genes (Massague 2000; von Bubnoff 2001). Hair
follicle morphogenesis is dependent upon the inhibition of BMP signaling as

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demonstrated by the observation that mice which overexpress Noggin have an
increased number of hair follicles (Plikus et al, 2004).

In this study, we set forth to examine the role of the BMP pathway during the
formation and growth of the mouse nipple. We demonstrate that BMP signaling is
central to a series of cell fate decisions altering the competence status in the nipple
epithelium that leads to the formation of hairs, sebaceous glands, and normal nipple
epithelium within the same nipple.

97

4.2 Methods and Materials
4.2.1 Animals
KRT14-Noggin mice were identified as described previously (Plikus et al, 2004).
Msx2 –/– mice were obtained as a gift from Dr. Robert Maxson (USC), backcrossed
onto a C57bl/6 background and then identified as previously described (Satokata et
al, 2000). All animals were

anaesthetized and their nipples removed

and processed
for staining or SEM as described below.
4.2.2 Histological Histochemical and Immunohistological Staining
4.2.2.1 H&E and Trichrome Staining
Following excision of the mouse nipples, the specimens were fixed in 4%
paraformaldehyde (PFA) in phosphate buffered saline (PBS) overnight at 4
o
C
followed by a dehydration series and embedding in paraffin wax. Sections of the
fourth inguinal nipple were cut longitudinally at 5 μm thick. Standard hematoxylin
and eosin (H&E) staining was performed for basic histological analysis.
4.2.2.2 Immunostaining Sections
Specimens were fixed, dehydrated, and embedded as described above. Sections of
the fourth inguinal nipple were stained with antibodies against Keratin 14 (KRT14)
(Lab Vision) and β-catenin (Sigma) to visualize signaling. Staining for pSMAD1,5,8
(Cell Signaling) was performed to visualize BMP signaling. Binding of the
secondary antibody and visualization using AEC (Vector) were done according
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to
the manufacturer’s protocol and as described in Jiang et al, 1998. Visualization and
photos were taken using the Trestle MedMicroscopy slide scanner fitted on an
Olympus BX51 microscope.
4.2.2.3 Staining of Sebocytes with Oil Red O
Oil Red O staining was modified and performed based on the protocol described
previously (Guha 2004). Briefly, 5-um frozen sections of the adult mouse nipple
were post-fixed in 4%PFA, washed with distilled water followed by two changes of
100% propylene glycol for 5 minutes each. The sections were stained for 7 minutes
with Oil Red O, followed by rinsing in 85% propylene glycol and then in distilled
water for 3 minutes each, and then counterstained with hematoxylin and mounted.
4.2.3 In situ hybridization
Section in-situ hybridization was performed as previously described but with some
modifications (Jiang et al, 1998). Proteinase K treatment was performed for 10
minutes and all washing steps were performed for one half the time as described for
whole mount in-situ hybridization. PTHrP probe was a kind gift from Dr. Marcel
Karperien and described previously (Karperien 1996). Shh probe was a kind gift
from Dr. Saverio Bellusci (USC).
4.2.4 Scanning Electron Microscopy
The specimens were excised and fixed overnight at 4
o
C in Karnovsky's fixative
(2% paraformaldehyde, 2% glutaraldehyde, 5mM CaCl
2
in 0.1M cacodylate
99

buffer, pH

7.4). After fixation, the specimens are rinsed several times with 0.1M cacodylate
buffer for a minimum of 15 minutes, followed by post fixation with 2% osmium
tetroxide in 0.1M cacodylate buffer for 2 hours in the hood. This is followed by
rinsing with 0.1 M cacodylate buffer for a minimum of 15 minutes followed by
dehydration of the specimens using a graded alcohol series (50% for 15 min, 70% for
15 min, 85% for 15 min, 95% for 15 min, and 2 changes of 100% for 5 min. each).
The samples are then chemically dried using a series of HMDS
(hexamethyldisilazane; 50% HMDS in ethyl alcohol for 15 minutes followed by
75%, 85%, 95% and 2 changes of 100%). After the final change of 100% HMDS, the
excess is removed and the samples are allowed to air-dry in a hood over night. The
samples are then mounted on aluminum stubs with adhesive tabs and sputter coated
for 3 minutes.

4.2.5 Semi Quantitative RT-PCR
A total of five nipples were excised from WT, KRT14-Noggin +/-, KRT14-Noggin
+/+, and Msx2 -/- mice. Tissues were then ground using a mortar and pestle in liquid
nitrogen and homogenized using the QIAShredder (Qiagen). RNA was extracted
using the RNeasy kit (Qiagen). Finally RNA was transcribed into cDNA using a
first-strand reverse transcriptase kit for RT-PCR (Invitrogen). We analyzed the
tissue for levels of Agouti, MC-1R, attractin, and MITF previously described
100

(Sharov et al, 2005). We also analyzed the tissues for levels of ectopic KRT14-
Noggin (Plikus

et al, 2004), endogenous Noggin sense -GAGCAAGAAGCTGAGGAGGA and
antisense -GTGAGGTGCACAGACTTGGA , BMPR1A sense -
TCTCAAGCAGGGGTCGTTAC and antisense -CGCCATTTACCCATCCATAC,
and L32 sense -AGAAGGTTCAAGGGCCAGAT and antisense –
CAGCTCCTTGACATTGTGGA as a control. PCR was performed using the BioRad
MJ mini as follows: An initial dissociation at 95°C for 3 minutes followed by 30
cycles of dissociation at 95°C for 30 seconds, annealing at 55-58°C for 30 seconds,
and extension at 72°C for 30 sec. A final extension time of 10 minutes was then
performed and PCR products were held at 4°C until run on a 1% agarose gel for
visualization of the bands.
4.3 Results
4.3.1 Overexpression of Noggin leads to the conversion of nipple epithelium
into competent hair follicle forming epithelium
Scanning electron microscopy (SEM) was performed to visualize the presence of
hair follicles formed within the nipples of Noggin expressing mice compared to
control and Msx2 -/- mice (Fig. 22 a-d). The images obtained from SEM allowed us
to count the number of follicles protruding from the nipples. The effects of KRT14-
Noggin showed a dose-dependence in hair follicle formation. Heterozygous
Noggin mice had an average of 10 +/- 4.83 hairs per nipple, while homozygous
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mice averaged 15.25 +/- 1.71 hairs per nipple. No hair follicles were observed in the
nipples of control or Msx2 -/- mice (Fig. 22 e).

Fig. 22 Scanning electron microscopy and quantitation of hair follicles from adult, 3
month old parous mice. Control and Msx2 -/- mice show no hair follicles within the
nipple, respectively (a, c). Noggin overexpressing mice show ectopic hair follicles
extruding from the nipples (b, d). A graph indicates the trend that increased hair
follicle numbers correlates with increasing Noggin expression (e). Red arrows
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point to a hair follicle growing within the nipple. Size bar = 200 um.

103

Since the nipple is part of a specialized ectodermal organ (mammary gland) derived
from epithelial - mesenchymal interactions, alterations in BMP pathway activity may
result in a change in the competence of the epithelium allowing for the formation of
hair follicles. After histological staining was performed we examined the nipples for
any morphological changes that had occurred. Noggin expressing mice showed hair
follicles and sebaceous glands within the nipple (Fig. 23 b,c) that were absent from
wild type nipples (Fig. 23 a). Since Msx2 is known to have a SMAD binding site in
its promoter region, we also examined the nipples of Msx2 -/- mice at these stages.
To examine whether Msx2 mediated this activity, we checked Msx2 -/- mice for hair
follicle formation in the nipple epithelium. H&E staining reveals that Msx2 -/- mice
also showed no hair follicle formation within the nipple similar to controls (Fig. 23
d). This data suggests that suppression of BMP activity by Noggin leads to a
conversion of nipple epithelial cells into different ectodermal organs including hair
follicles and sebaceous glands. This activity does not appear to be mediated via
Msx2. Oil Red O staining was performed on frozen sections of the mouse nipples to
confirm the presence of sebaceous glands within the nipples of the transgenic
Noggin mice. Results show that Noggin acts in a dose dependent manner that
increases the number of ectopic sebaceous glands formed within the nipple (Fig. 23
f,g). WT and Msx2 -/- mice show no formation of sebaceous glands within the nipple
(Fig. 23 e, h, respectively). KRT14 staining was performed to show the location of
the transgene in KRT14-Noggin mice (Fig. 23 j, k) and stain the newly formed hair

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follicles in the Noggin mice. KRT14 staining was not altered in the transgenic mice
compared to controls (Fig. 23 i-l).
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Fig. 23 Histological and immunohistochemical analysis of adult, parous 3 month old
wild-type and transgenic mouse nipples with decreased BMP activity. H&E stained
sections of control (a), KRT14-Noggin +/- (b), KRT14-Noggin +/+ (c), and Msx2 -/-
mice (d). Oil Red O staining to indicate the presence of sebaceous glands in the
Noggin overexpressing mice compared to Msx2 -/- and control mice (e-h). KRT14
immunostaining showing no change in nipple epithelium from control, Noggin over
expressing transgenic or Msx2 -/- mice (i-l) but increased epithelial expression
surrounding ectopic hair follicles in Noggin overexpressing mice (j, k). Black arrows
point to pilosebaceous units. Size bar = 200 um.

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4.3.2 Overexpression of Noggin leads to ectopic expression of Shh while
suppressing PTHrP
We were interested in ascertaining the molecular pathways affected as a result of
suppressing BMP pathway activity. In-situ hybridization results reveal that Noggin
overexpression decreases the expression levels of PTHrP compared to controls (Fig.
24 a-c). PTHrP does not appear to be affected in the Msx2 -/- mice (Fig. 24 d). Shh
was also shown to be expressed in the epithelium of the ectopically formed hair
follicles in the nipple compared to control and Msx2 -/- mice (Fig. 24 e-h). This
alteration in signaling within the basal layer of the epidermis causes the expression
levels of PTHrP and Shh to be altered. The nipples of Msx2 -/- mice however do not
have hair follicles present suggesting that Msx2 is not involved in setting up the
formation of a competent epithelium that allows for hair follicles to form in the
nipple. This series of molecular events leads to the formation of a competent nipple
sheath and allows for the formation of hair follicles in the nipple of KRT14-Noggin
mice. We demonstrate that inhibiting BMP signaling within the basal keratinocytes
is permissive for an epithelial fate change that allows for the formation of hair
follicles in the mouse nipple where hair normally would not form.
107

Fig. 24 Expression patterns for PTHrP, Shh, and pSmad1/5/8 in the nipples of adult,
parous 3 month old WT, Noggin overexpressing, and Msx2 -/- mice. PTHrP
expression appears to decrease in a dose dependent fashion in Noggin
overexpressing mice compared to WT and Msx2 -/- mice (a-d). Noggin
overexpressing mice show an increase in epithelial Shh expression surrounding the
ectopic hair follicles but no change in nipple epithelial expression compared to
control and Msx2 -/- mice (e-h). p-Smad1/5/8 immunostaining is reduced in the
nipple epithelium and increased in the epithelium surrounding the ectopic hair
follicles in Noggin overexpressing mice compared to WT and Msx2 -/- mice (i-l).
Red arrows in panels b and c indicate areas with reduced PTHrP expression while
black arrows in panels f, g, j, and k point to increased Shh surrounding ectopic hair
follicles. Size bar = 200 um.

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4.3.3 Overexpression of Noggin leads to pigmented nipples
To understand the role of the BMP pathway in mouse nipple development, control
and transgenic mice overexpressing Noggin under the KRT14 promoter were
compared visually, microscopically and at the molecular level. For this purpose,
mice were shaved prior to macroscopic observations (Fig. 25 a-d). Upon visual
inspection the nipples of the KRT14-Noggin +/- and KRT14-Noggin +/+ mice
appeared pigmented compared to wild type controls (Fig. 25 e-g). Nipple
pigmentation was also increased in Msx2 -/- mice compared to controls (Fig. 25 h).
Pigmentation is clearly evident from hematoxylin stained sections of the transgenic
Noggin mice and the Msx2 -/- mice compared to controls (Fig. 25 i-l). Since Msx2
has a BMP response element in its promoter it is possible that BMP is acting through
Msx2 to suppress pigmentation. This suggests that Msx2 is likely involved in the
cessation of pigment production or in the maintenance of pigmentation within the
mouse nipple since pigmentation persists beyond puberty in ectopic Noggin
expressing and Msx2 -/- mice compared to wild-type mice.
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Fig. 25 Nipple Pigmentation. Gross view, higher magnification of gross view, and
hematoxylin staining of 8 month old mouse nipples were examined in WT (a, e, i)
KRT14-Noggin +/- (b, f, j), KRT14-Noggin +/+ (c, g, k) and Msx2 -/- (d, h, l) mice.
Note pigmentation is present in the KRT14-Noggin overexpressing and Msx2 -/- but
not wild type mice. The dark patches represent hair growth in the anagen phase of
the hair cycle (d). Red arrows point to pigmentation. Size bar for panels e-h = 500
um, for panels i-l = 200 um.

110

BMP signaling has been shown to control the expression of Agouti protein which
links the BMP and melanocortin-1 receptor (MC-1R) signaling pathways during the
production of pigment

in the hair follicle (107 Sharov,A.A. 2005). We first explored
the levels of Agouti in our mice. As predicted the levels of Agouti increase in a dose
dependent manner as a result of Noggin overexpression but remain unchanged in
Msx2 -/- mice. To further explore the mechanisms involved in the continued
production of pigmentation in the nipples of adult KRT14-Noggin overexpressing
and Msx2 -/- mice, the expression

of the components of the MC-1R signaling
pathway including attractin, MC-1R, and melanocyte-restricted microphthalmia-
associated transcription factor M-MITF, a regulator of melanogenesis, were
compared to controls. We also compared the levels of BMPR1a since studies suggest
that BMPR1a may be upregulated causing an increase in mesenchymal signaling
leading to the increased BMP signaling in the epithelium. Results from semi-
quantitative RT-PCR indicate that in fact the MC-1R pathway is activated in the
nipples of Noggin overexpressing mice as well as being partially activated in Msx2 -
/- mice compared to controls. M-MITF is upregulated in the nipples of Noggin
overexpressing and Msx2 -/- mice. Attractin levels appear to be decreased in the
nipples of Msx2 -/- mice compared to all others while MC-1R itself appears to be
only slightly upregulated in KRT14-Nog +/+ mice compared to the others. BMPR1a
expression has not been altered in the nipples as a result of decreasing BMP activity

111

in both Noggin over expressing and Msx2 -/- mice (Fig. 26). Control gene (L32)
expression was consistent throughout the samples.
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Fig. 26 Expression of the melanogenesis regulators in nipples of 8 month old WT,
Noggin overexpressing, and Msx2 -/- mice. Nipples were harvested from WT and
transgenic mice and analyzed for Agouti, MC-1R, attractin, and M-MITF expression
by RT-PCR. Levels of endogenous levels of Noggin, ectopic levels of KRT14-
Noggin, BMPR1A, and control L32 genes were also compared. Compared to
controls Noggin over expressing mice have increased expression of ectopic Noggin,
Agouti, and M-MITF; Msx-2 -/- mice have increased M-MITF and decreased
Attractin.
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4.4 Discussion
4.4.1 Signaling pathways involved in hair follicle and nipple formation
Human nipples are often pigmented structures that include melanocytes located in
the basal layer of sebaceous glands that are not associated with hair follicles
(Montagna 1970). Our data examined the role of Noggin in murine nipple and hair
follicle morphogenesis. The transgene is clearly misexpressed in the entire basal
layer of the epithelium in these animals as previously reported (Plikus et al, 2004).
Ectopic pilosebaceous unit formation including hair and sebaceous glands is another
phenotype of these KRT14-Noggin transgenic mice. This suggests that inhibition of
BMPs likely drives keratinocytes toward hair follicle fate determination in the skin
while suppressing the nipple epithelium differentiation pathway. Lineage selection
by the common progenitor cells in the skin is therefore dependent upon the BMP
signaling pathway. Recent studies have shown that removal of Shh signaling in the
epithelium secondarily causes increased levels in the mesenchyme contributing to
induction of hair follicles (Gritli-Linde 2007). Our results indicate that noggin
overexpressing mice exhibit epithelial Shh expression in the abnormal hair follicles
which form within the nipple. Shh expression is not altered in the overlying nipple
epithelium suggesting that the increase in Shh expression is secondary to the
formation of the hair follicles in the nipple.

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PTHrP signaling is required for the proper formation of the nipple. In the absence of
PTHrP signaling, mammary mesenchyme fails to form and in turn does not direct the
formation of nipple sheath in the overlying epidermis (Wysolmerski 1998; Dunbar
1999; Foley et al, 2001). Conversely, KRT14-PTHrP mice overexpressing PTHrP in
the basal keratinocytes causes the conversion of subepidermal mesenchyme into
condensed mammary mesenchyme (Foley et al, 2001). This leads to the suppression
of hair follicle development and causes the entire ventral epidermis between the
mammary lines to acquire characteristics of the nipple sheath. Interestingly,
overexpressing PTHrP in this manner has been suggested to lead to the accumulation
of dermal melanocytes and thus there is pigmentation of the ventral skin between the
mammary lines of the KRT14-PTHrP mice (Abdalkhani 2002). A recent paper
confirms our finding that PTHrP is directly affected by altering BMP pathway
activity (Hens et al, 2007), but these authors show that PTHrP regulates BMP
receptor expression. In contrast, our data shows that BMP can regulate PTHrP
activity since it is down regulated in Noggin transgenic mice. Perhaps there is a
feedback loop through which BMP and PTHrP pathway activity is regulated.
Furthermore, our data suggests that perhaps both PTHrP and BMP pathway signaling
play a role in the production of pigmentation. However, this pigmentation was not
maintained in the KRT14-PTHrP adult mice, suggesting that BMP but not PTHrP is
required for the maintenance of pigmentation produced by these dermal melanocytes.

115

One way that BMPs can exert their effects is through Msx1 and Msx2 which are
BMP-responsive homeodomain-containing transcription factors that have been
shown to participate in the epithelial - mesenchymal signaling during development
(Phippard et al, 1996; Satoh 2004; Satokata et al, 2000). Both Msx1 and Msx2 are
important to the normal development of epithelial appendages including hair and
mammary glands. In mice that are deficient for Msx1 and Msx2, mammary
development fails at the placode stage (Satokata et al, 2000). Msx2 alone has also
been shown to be expressed within the dense mammary mesenchyme (Phippard et al,
1996; Satokata et al, 2000). Satokata (et al, 2000) reported that mammary buds form
in Msx2 -/- mice but their development subsequently arrests at E16.5 and no ductal
outgrowth is formed (Satokata et al, 2000). While Msx2 was previously reported to
be required for the outgrowth of the mammary bud in mice (Satokata et al, 2000), we
have found that these mice, in our hands, can form normal and functional mammary
ducts and nipples. Recently Hens (et al, 2007) also reported that these mice are
capable of forming normal mammary buds (Hens et al, 2007). Hens et al, 2007 report
that the ability of PTHrP to suppress hair follicle development is dependent upon
Msx2 (Hens et al, 2007). They report that mesenchymal expression of Msx2 is able
to interfere with the ability of hair follicle formation in the epidermis surrounding the
nipple. But this observation is based on epithelial cells that are converted to nipple-
like epithelium via the over expression of PTHrP. In contrast, our observations are
based directly on nipple epithelium. BMPs regulate Msx expression by virtue of a
BMP responsive region in their promoter (Brugger et al, 2004). Our Msx2 -/-
116

adult mice display normal formation of mammary glands (data not shown) and
pigmented but hairless nipples. These mice are capable of breeding and nursing their
pups. This indicates that Msx2, or rather the lack of Msx2 is required for the
maintenance of pigmentation within the nipple but is not involved in the lateral
inhibition of hair follicle formation within the developing nipple.

Semi-quantitative RT-PCR indicates that BMPR1a expression has not been altered in
the nipples as a result of decreasing BMP activity in both Noggin over expressing
and Msx2 -/- mice. This tells us that the BMP receptor is not upregulated as a result
of increasing Noggin levels or decreasing Msx2 levels. MC-1R pathway activity has
been altered in the transgenic mice involved in this study. In all mice that exhibit
pigmentation, M-MITF is expressed in the nipples. This suggests that the
maintenance of pigmentation is due to an increase in MC-1R pathway activation.

In this study we demonstrate that overexpression of Noggin under the KRT14
promotor is sufficient to alter the competence of nipple epithelium. Here we propose
a model of molecular activity (Fig. 27). By overexpressing Noggin in the epidermis
we subsequently decrease BMP pathway activity as well as PTHrP expression. This
leads to a conversion of nipple epithelium to follicular epithelium containing hairs.
Levels of Shh in the nipple epithelium of Noggin overexpressing mice were
unchanged from controls while Shh is expressed within the ectopically formed hair
follicles preventing reversion back to nipple epithelium. Overexpression of
117

Noggin in addition to loss of Msx2 leads to an increase in pigmentation found within
the nipple. This pigmentation is maintained due to an increase in MC-1R pathway
activity. Previous studies show that the epidermis contains a population of stem cells
that can proliferate and give rise to differentiating daughter cells that signal to the
mesenchyme to form the various ectodermal organs including mammary glands, hair
follicles, and sebaceous glands (Niemann 2002). Our study brings out the possibility
that the epithelial stem cells in the mammary gland and nipple region can be altered
to produce the formation of hair instead of hairless nipple epithelium. We have yet to
determine whether the altered nipple epithelium is capable of reverting back to
hairless, unpigmented nipple epithelium.
118

Fig. 27 Effects of BMP signaling on epithelial appendages (a). Model of normal
BMP signaling on cell fate within the mouse nipple (b).

119

4.4.2 Evolution of the nipple and mammary gland
The evolution of the nipple and mammary gland is interesting and of particular
importance since the development of these structures allows for improved milk
delivery to the young and thus increased survival. Many species have evolved ways
to feed their young with varying success. Interestingly eggs have been proposed to
be an important element in mammary gland evolution (Oftedal 2002a; Oftedal
2002b). Early Synapsids, are believed to have used secretions from their glandular
skin to replenish fluids to the egg shells surrounding their young, as do current
amphibians such as salamanders (Widelitz et al, 2007). The secretions of
Monotremes, on the other hand, trickled from mammary patches consisting of
nippleless areolae-like skin, to a hairy coat in order to feed their young. Today’s
platypus uses specialized “mammary hairs” located along two presumptive
“mammary” lines which run along their ventral surface (Widelitz et al, 2007). Since
altricial newborns rely more upon early feedings from their mothers than their
precocial counterparts, a hairless areola with a nipple was developed as an efficient
means to feed the young. Mice do not exhibit an areolar complex as humans do but
merely contain nipple epithelium adjacent to normal skin. Therefore, we can
speculate that the evolution of mammary glands and nipples may be associated not
only with the live birth of mammals but also with the molecular gain of BMP
pathway activity. This would allow for specialized mammary glands and nipples to
form while preventing hair formation in this region.

120

5.0 Chapter 5 – Discussion
Using two animal model systems, I explored the role of hormones in regulating the
growth of skin appendages (ectodermal organs). In general in humans it is thought
that hormone stimulation has the ability to increase the surface area of skin
appendages. Clearly mammary gland development through puberty, pregnancy, and
parturition are affected by levels of sex hormones. To better understand how sex
hormones regulate growth control, I turned to another skin appendage model,
feathers. Based upon the literature, the understanding is that estrogens are
responsible for the phenotype of the female feather. In the case of chicken rectrices,
androgens had surprisingly little effect, yet estrogens reduced the growth rate and
duration of growth. Utilizing current methodologies we were able to show that the
female feather is a result of the local conversion of estrogen. Aromatase is expressed
in the epithelium of the female to convert testosterone into estradiol, while the lack
of 5a-reductase further ensures that any testosterone present in the female feather is
converted to estradiol. This over compensatory expression of molecules ensures the
female phenotype and is necessary since the default sex and phenotype of the
chicken is male. Further studies need to be performed to verify the location and cell
types of the enzymes and receptors. This can easily be accomplished by performing
in-situ hybridization for the molecules involved. Sex hormone dependent epithelial
growth has been shown in the case of the chicken feather to be dependent on the
local estrogen / androgen ratio, integrating hormones produced systemically and
locally in the skin. Through this study we were able to demonstrate that local
121

hormone metabolism may produce sexual dimorphisms. This can have broad
implications for the treatment of hormone-dependent cancers.

We hypothesized that the BMP pathway is required for normal mammary gland
formation and in the absence of increased levels of estrogen, Noggin promotes
abnormal stroma formation leading to increased epithelial cell density. KRT14-
Noggin mice display dilated ducts and delayed involution compared to wild-type
controls. Upon close inspection collagen I was present surrounding the ducts and
milk products remain inside. This suggests that the BMP pathway is required for the
proper involution of the mammary gland and the proper organization of the stroma
surrounding the ducts. Further experiments are needed to show how inhibition of
BMP signaling induces increased collagen accumulation. This can be addressed by
examining SMAD binding sites in the collagen promoter via ChIP assays. Also it is
important to understand whether the collagen remaining around the ducts in the
KRT14-Noggin mammary glands is due to increased synthesis or decreased
degradation. This can be determined by staining for MMPs which are known to be
involved in matrix degradation. Should MMPs be reduced compared to wild-type
then the answer is simple, the collagen present is due to decreased degradation.

During evolution the BMP pathway was most likely gained to ensure the proper
cycling of the mammary gland following parturition. The gain of this pathway was
also involved in the development of the nipple. KRT14-Noggin mice exhibit the
122

formation of hair follicles within the nipple. The presence of hair creates difficulty
feeding their young. This hair formation is due in part to the reduced expression of
pSMAD and PTHrP. Future studies need to be performed to show exactly how
BMPs regulate the expression of PTHrP. This can be accomplished by utilizing the
ChIP assay to identify if in fact downstream members of the BMP pathway are
capable of binding the promoter and activating transcription of PTHrP. Another
method is to use PTHrP reporter constructs, transfected into cells, and activated by
BMP protein to test whether BMP is capable of activating expression of the
contructs. In our mouse nipple and mammary gland model we found that the BMP
pathway plays a major role in mediating hormone response. BMP may be required
at the end of the pregnancy cycle to prepare the tissue for a new cycle. Down
regulation or disruption of the BMP pathway leads to increased extracellular matrix
accumulation which has been associated with increased breast density. Increased
breast density is a high risk factor for breast cancer.

While this research is basic in nature, the work presented a unique opportunity to
enhance our understanding of the molecular mechanism of how the sex hormone
pathway is linked to a growth related pathway in a physiological context. It should
be noted that studies in Drosophila or C. elegans have brought major new
understanding to cancer biology. In fact beta catenin (β-cat), wnt, and sonic
hedgehog (Shh) were all first discovered using the Drosophila model. I hope that our
novel experimental models contribute to further understanding of the hormone
123

dependent growth of breast and prostate cancer, which are clinically more important
but less susceptible to experimental analyses.
124

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Abstract (if available)

Abstract The mammary gland, hairs, feathers, etc. are all skin appendages derived from epidermis as a result of epithelial - mesenchymal interactions. The growth of these epithelial organs shares fundamental morphogens including BMP, SHH, Wnt/beta catenin, etc. Mutation or de-regulation of these morphogens have been identified in many human diseases including tumors and genetic diseases. Sex hormones also play a major role in regulating the growth and phenotypes of these epithelial organs (sexual dimorphism). How the sex hormones are coupled to the morphogens is mostly unknown. In this thesis I utilized two distinct epithelial appendage models, a chicken tail feather (between rooster and hen), and the mouse mammary gland model. In the chicken tail feather model, we were able to show that the female feather phenotype is dependent on the local conversion of estrogen. We also utilized the transgenic KRT14-Noggin (BMP antagonist) mouse that shows abnormal mammary gland stroma formation and delayed involution. Imbalance of BMP pathway lead to increased collagen I deposition surrounding the ducts and hair formation within the nipple of these transgenic mice. Taken together, this opens the possibility that the sex hormone pathway may be involved with the BMP pathway. We offer novel experimental models that may contribute to further understanding of the hormone dependent growth of breast and prostate cancer, which are clinically more important but less susceptible to experimental analyses.

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Asset Metadata

Creator Mayer, Julie Ann (author)

Core Title Skin appendage growth control by hormones and morphogens

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Pathobiology

Publication Date 12/08/2009

Defense Date 07/17/2007

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag androgen,BMP pathway,Estrogen,feather,hair,mammary gland,nipple,Noggin,OAI-PMH Harvest,receptor

Language English

Advisor Widelitz, Randall (committee chair), Chuong, Cheng-Ming (committee member), Pike, Malcolm C. (committee member), Press, Michael (committee member)

Creator Email juliemay@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m972

Unique identifier UC1425299

Identifier etd-Mayer-20071208 (filename),usctheses-m40 (legacy collection record id),usctheses-c127-595875 (legacy record id),usctheses-m972 (legacy record id)

Legacy Identifier etd-Mayer-20071208.pdf

Dmrecord 595875

Document Type Dissertation

Rights Mayer, Julie Ann

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Repository Name Libraries, University of Southern California

Repository Location Los Angeles, California

Repository Email cisadmin@lib.usc.edu