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Morphoregulation and cycling of ectodermal organs

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Morphoregulation and cycling of ectodermal organs

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MORPHOREGULATION AND CYCLING OF ECTODERMAL ORGANS

by

Maksim V. Plikus

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)

May 2007

Copyright 2007 Maksim V. Plikus

ii
Dedication
I would like to dedicate this work to my teachers and mentors: Dr. Cheng-Ming
Chuong and Dr. Alex Astrowski without whom my research will not be possible,
and to my parents who always believed in me.

iii
Acknowledgments
I would also like to acknowledge Dr. Randall Widelitz, Dr. Wen Pin Wang, Dr.
Ting-Xin Jiang, Dr. Robert Maxson, Dr. John Suundberg, Dr. Ralf Paus, Dr. Ruth
Baker, Dr. Philip Maini, Dr. Maggie Zeichner-David, Dr. Julia Reyna, Dr. Pablo
Bringas, Dr. J. G. M. Thewissen, DR. Malcolm L. Snead, Dr. Yang Chai, Zina
Zhang, Julie Mayer, Lisa Doumak, and all past and present members of Dr.
Chuongs lab.

iv
Table of Contents
Dedication ii
Acknowledgements iii
Abstract v
Introduction 1
Chapter 1: Morphoregulation of ectodermal organs 109
Chapter 2: Morphoregulation of teeth 156
Chapter 3: BMP activity in spreading of hair cycle regenerative waves 208
Conclusion 260
References 264

v
Abstract
Ectodermal organs are produced through the series of epithelial-mesenchymal
interactions followed by topological transformation of the otherwise flat
ectodermal layer. Fundamental signaling pathways regulate development of
ectodermal organs. Fine tuning of the organs size and shape during development
is achieved by mild spatial-temporal changes of signaling pathway activity rather
than on-off switches. In my work I have investigated the extent of
morphoregulatory activities of Bmp signaling pathway. I have modulated normal
Bmp signaling by overexpressing of Bmp antagonist noggin under keratin 14
promoter in K14-noggin mice. I have shown that changes in Bmp pathway
activity can alter various ectodermal organs at different developmental stages:
induction (increased number of pelage hair follicles, formation of compound
vibrissa follicles, claw agenesis, transdifferentiation of sweat glands into hair
follicles etc.), morphogenesis (defect of eyelids opening, enlargement of external
genitalia etc.) or differentiation (incomplete differentiation of claw plates,
retarded differentiation of hairy spines). I have further shown that in
morphologically complex teeth, Bmp pathway regulates all of the above stages of
development.
Furthermore, I have identified a new physiological mechanism of hair cycle
control. I have shown that rather than being an autonomous intra-follicular event,
the critical telogen-anagen transition during the hair cycle is largely regulated by
signals originating from the extra-follicular environment and neighboring hair
follicles. I have conclusively identified skin-wide changes in Bmp signaling
activity as the major regulatory factor of telogen-anagen transition in large

vi
populations of hair follicles. My findings allow to view the hair cycle in the
content of the microenvironment surrounding the hair follicle. They allow us to
explain how hair follicles make decisions on hair cycle progression by listening
to other follicles and inter-follicular signaling centers and how multiple hair
follicles can coordinate their hair cycles to form macroscopic growth patterns.
My findings open new area for hair cycle control studies that takes an integrative
approach and views hair follicles in relation to the surrounding stroma and
neighboring follicles. My work proves morphoregulation hypothesis by showing
how mild spatial-temporal changes in a single fundamental signaling pathway
can modulate all stages of development of multiple ectodermal organs.

1
Introduction
In this introductory chapter I discuss the current status of knowledge in the field
of developmental biology of mouse skin ectodermal organs. I compile many
representative figures from the explosively increasing literature.

1.1 Skin ectodermal organs – model system in developmental biology
The skin forms the interface between the organism and environment. For species
to adapt successfully, the skin has evolved specific ectodermal organs in different
regions for temperature homeostasis, defense, sensory, communication, breeding,
etc. The laboratory mouse has become a rich resource to learn how these
ectodermal organs are made, maintained, repaired, and regenerated. Here I survey
pelage hairs, vibrissae, sebaceous glands, sweat glands, nails, volar pads,
mammary glands, etc. For each ectodermal organ, I first describe the morphology
and structure, followed by developmental stages and involved molecular
signaling pathways. While there are different morphologies and functions, these
ectodermal organs all result from epithelial-mesenchymal interactions and share
developmental phases of induction, morphogenesis, differentiation, and cycling
(only in some organs). Similar signaling molecular "tools" including, Eda, Shh,
Bmp, Fgf, Notch, Msx, etc. are used repetitively in different developmental
phases of the same organ and in different organs, leading to different
consequences. Since the skin is the most apparent organ, many interesting
findings involving the skin ectodermal organs, some expected and some
unexpected, continue to emerge when genetically engineered mice are made. As a
result, rapid progress has been made that helps make skin biology a mainstream

2
of research into understanding signaling pathways in organogenesis. Hair
research also has become a frontier of stem cell biology / regenerative medicine.
The skin forms the outermost boundary of all higher organisms and is in direct
contact with the outside world. Often animals have to survive and function under
extreme temperatures, humidity, solar radiation, mechanical abrasion, etc. The
skin enables animals to cope with all of these stresses and many others.
Environmental conditions are dynamic and can change with annual seasonal
cycles or shift over millions of years on a geological time scale. During the
process of evolution, mammalian skin successfully adopted to withstand dramatic
environmental changes. Key strategies for adaptation include transformation of
the flat skin epithelium into various types of skin appendages with different
levels of complexity to perform specialized tasks with maximum efficiency.
While primarily functioning to preserve homeostasis, the skin also evolved to
serve functions in communication, mechano-sensory evaluation of the
environment, defense, attack, mimicry, etc. (Chuong edit 1998; Chuong CM et
al., 2002). Here we focus on the skin of the laboratory mouse because this animal
is the major experimental model for biomedical research representing mammals.
Skin is the product of epithelial-mesenchymal interactions followed by
topological transformation of the otherwise flat epidermal layer into different
types of appendages (Figure 1.1). For each type of skin appendage we will first
describe its morphology, physiology, and development, then summarize known
epithelial-mesenchymal interactions important for morphogenesis, and review
molecular signaling controls in development into the adult anatomic structures.
Examples of spontaneous mutant mice and genetically engineered mice will be

3
used to illustrate these processes. We provide here an overview of mouse skin
ranging from topological arrangements, regional specificity, size regulation,
functional morphology, etc. Developmental changes demonstrate how each
distinct ectodermal organ is transformed from an initial flat piece of ectoderm
during embryogenesis into complex anatomic structures. Integration of the
current understanding of the genes and their protein products that are critical in
embryogenesis, maintenance, and normal cycling are integrated throught the
discussions. In the last decade, there was rapid progress in the field of mouse skin
research with an explosive accumulation of knowledge that changed this field
from superficial "skin deep" research into mainstream molecular biology with the
skin serving as an organ of focus for development and more important complete
regeneration in the case of the hair follicle cycle. We can expect that mouse skin
will continue to be the primary window for understanding molecular
organogenesis. However, at this stage, much of our understanding of molecular
pathways is at the level of "X molecular pathway involved". Namely, when the
homeostasis of that molecule is perturbed, the formation of that ectodermal organ
is disrupted. We continue to learn that often several molecules are involved. Yet,
we do not know how to put these molecular pathways together to engineer an
ectodermal organ. This knowledge is critical for the progress of stem cell biology.
In the near future, we may have enough stem cell populations (whether
embryonic or adult stem cells) to learn how to induce them to differentiate with
certain factors in vitro, but we still need to learn how to guide stem cells to form
tissues and organs with the right three dimensional topology (Chuong et al., in
press). Skin ectodermal organs are likely to be the models for these trial.

4
Figure 1.1 Diversity of skin appendages in mouse. Schematic drawing highlights
the topics covered

5
The laboratory mouse continues to be the major model for diseases both as
natural or spontaneous mutant mice or those produced by a variety of genetic
engineering methodologies (Sundberg and Ichiki 2005). Examples of the types of
diseases we now have good models for include different types of alopecia,
epidermolysis bullosa, inflammatory diseases, and many others (Sundberg and
King, 2000). In addition to being used to further our understanding of basic
biology, mutant laboratory mice can also be used as models for new treatments
for specific diseases. The accessibility of the skin makes it a primary first line
target.

Finally, when we learn significantly about the laboratory mouse, it is also
valuable to apply the knowledge to the wild mouse. The comparative knowledge
of molecular biology of the skin ectodermal organs between laboratory mouse
skin and diverse rodents in nature will help us to gain new perspectives in the
evolution of diverse skin ectodermal organs.

1.2 Epidermis
Skin is made of epidermis and dermis. Dermis consists of fibroblasts and
extra-cellular matrix which are important for the physical properties of the skin as
well as biochemical interaction with the epidermis. In different parts of the body,
different types of skins (e.g., thin skin versus the volar pad) and different types of
skin ectodermal organ can form (e.g., hairs, glands), probably specified by the
dermis. In addition, the dermis is composed of vasculatures, nerves, muscle,

6
adipose tissues, etc. Although the dermis is obviously important, in this chapter,
we will focus on the epidermis in skin ectodermal organs.

1.2.1 Morphology
The epidermis is a cornified stratified epithlium covering the skin surface, from
which the follicles extend into the dermis. Mouse skin epidermis is significantly
thinner than human. This is because mouse integument is dominantly covered by
the fur coat, which contributes to the protective role of the epidermis. In
nonhaired skin areas, such as the nasal planum, the foot pads, the epidermis is
markedly thicker and exhibits rete pegs. This topic has been frequently reviewed
elsewhere (Sundberg et al., 1996; Sundberg and King, 2000) and will be briefly
summarized in this section.

Five distinct layers are identified in the epidermis (Eckert RL et. al., 1997). Cells
of the innermost basal layer (stratum basalis) are relatively undifferentiated and
have high proliferatory activity that ceases as they migrate into the suprabasal
layers of the epidermis. Basal cells sit on the basal lamina and are anchored to it
via hemidesmosomal junctions. Upon entering the suprabasal layers these
epidermal keratinocytes begin a program of terminal differentiation. The switch
from proliferation to differentiation is accomplished by changes in the expression
profile of keratins and other terminal differentiation markers such as envoplakin
(Evpl) and periplakin (Ppl; Ruhrberg et. al., 1996; Ruhrberg et. al., 1997),
involucrin (Ivl; Rice and Green, 1979; Nemes et. al., 1999), transglutaminase 1
(Tgm1; Kim et. al., 1995), members of small proline rich-like (Sprrl) family.

7
While basal cells produce keratin 5 and keratin 14, suprabasal keratinocytes start
to express keratin 1 and keratin 10. The spinous layer (stratum spinosum) is the
first suprabasal layer and it is characterized by an abundance of desmosomes and
by the onset of involucrin production (Upon rise of Ca
2+
concentrations, Tgm1
crosslinks Ivl molecules with each other to form two-dimentional oligomeric
mesh. It also cross-links involucrin to envoplakin. This initiates assembly of the
cell envelope; Steinert and

Marekov 1999). Next is the granular layer (stratum
granulosum). Cells of the granular layer have distinct basophilic granules
referred to as keratohyalin grabules - that contain loricrin, filaggrin, hornicrin,
cystatin-alpha, and lipids. As cells enter the stratum lucidum layer (transition
zone) they undergo programmed cell death, the keratin intermediate filament
network undergoes final assembly and the cornified envelope forms. These
components are important for the envelope formation of the stratum corneum
which is crucial in the defense of the individual. As this chapter focuses more on
ectodermal organ formation, readers who want more information on this topic
should be refereed to Elias, 2005. Upon completion of this maturation process,
already dead cornified cells (corneocytes) become part of the uppermost stratum
corneum (layer 5). The stratum corneum has an elaborate organization of
intercalated flattened corneocytes embedded in the lipid matrix. The stratum
corneum functions as the primary mechanical barrier between the animal and its
environment to prevent desiccation, toxin entry, and microbial infection (Eckert
RL et. al., 1997). The stratum corneum maintains a status of equilibrium between
continuous shedding and replenishment by constant production and

8
differentiation of underlying layers thereby allowing it to keep the skin surface
intact (Byrne et. al., 2003).

1.2.2 Developmental and molecular signaling
Although the epidermis appears to have a simple morphology of stratified
epithelium in comparison with the elaborate organization of its appendicular
derivatives (HFs in particular), the process of the epidermis formation is complex
and involves several key control mechanisms. There is a large amount of
literature covering this topic and molecular pathways including adhesion
molecules, cytoskeleton, signaling molecules, protein kinases, and transcription
factors are involved (Fuchs and Raghavan, 2002; Watt FM, 2002; Bikle DD et
al., 2001; Denning MF, 2004; Lechler T and Fuchs E, 2005). Here we only give
couple examples. During embryonic development epidermis progreses form
single-layered to stratified morphology. Transition between these two stages from
E15.5 onwards involves asymmetric division of the basal epidermal cells that
gives rise to committed suprabasal cells that move upwards (Lechler T and Fuchs
E, 2005). It was further shown that launch of the asymmetric division and
stratification is achived through the control of the mitotic spindle orientation
within dividing basal epidermal cells. Apical (opposite to basal membrane)
accumulation of LGN−mInsc−Par3 crescent complex (LGN (mammalian Partner
of Inscuteable (Pins) analog), mInsc (mouse homolog of Inscuteable) and Par3)
and its interaction with NuMa protein regulates perpendicular polarization of
mitotic spindle (Lechler T and Fuchs E, 2005; Koster MI and Roop DR, 2005).

9
1.2.2.1 c-Myc signaling
c-Myc (see alos section IV.C) was shown to play a complex role in maintaining
the proper balance between epidermal proliferation and differentiation.
Overexpression of human MYC2 under the control of the loricrin promoter in
ML-MYC2 mice results in inhibition of terminal differentiation of keratinocytes
characterized by an expanded suprabasilar proliferating domain (Waikel RL at.
al., 1999). It was further shown that c-Myc also regulates the maintenance of
epidermal stem cells in skin. Overexpression of MYC2 in the epidermal stem cell
compartment under the KRT14 promoter in K14-MYC2 mice caused dysregulated
proliferation and rapid depletion of the epidermal stem cells. As a consequence
adult K14-MYC2 mice also develop spontaneous skin erosions. It was suggested
that the stem cell depletion effect of c-Myc is mediated through the
downregulation of Itgb1 (Integrin beta 1) expression, which by itself is essential
for stem cell maintenance (Gandarillas A and Watt FM, 1997; Waikel RL, 2001).

1.2.2.2 p63 signaling
During the normal development of epidermis, p63 is expressed in the ectoderm
prior to its stratification and later p63 expression restricts to the basal layer of the
already stratified epidermis (Koster MI et. al., 2004). Transgenic mice lacking
p63 (p63
Brdm1
/ p63
Brdm1
and p63
Brdm2
/ p63
Brdm2
) fail to develop stratified
epithelia and epithelial appendages (Mills AA et. al., 1999; Yang A et. al., 1999).
In these mice LGN−mInsc−Par3 crescent complex does not show apical
accumulation and perpendicular orientation of the mitotic spindle does not occurs
(Lechler T and Fuchs E, 2005). Lack of a skin barrier due to the absence of

10
epidermal stratification causes quick dehydration and death just hours after birth
of these mutant mice. It was further shown that p63 is additionally required for
maintenance of the proliferative potential of basal keratinocytes

in mature
epidermis (Koster MI et. al., 2004). During evolution of the mammals, the basal
layer of the epidermis interact with dermis and gave rise to different types of skin
ectodermal organs important for adapting animals to different niches (Chuong,
1998). In the following sections, we describe the morphogenesis of these
ectodermal organs and some molecular signalings begun to be revealed. Many of
these signals, particularly those derived from dermis and related to regional
specificity, remain to be unknown challenges.

1.3 Hair Follicles
Body hairs are the most dominant feature when one first sees a mouse. On the
mouse, one can only find what appears to be glabrous (naked) skin on the foot
pad and behind the ears. Hair follicle (HF) morphogenesis and cycling have been
important research models for cell and developmental biologists in recent years
for many reasons. The skin is the most obvious place to find changes in
phenotypes due to spontaneous or genetically induced mutations. HFs have
robust regenerative power where an entire organ can form and reform from organ
specific stem cells on a regular and predictable basis. Recent advances in the field
of stem cell research found that HF stem cells contribute to the production of
extra-follicular tissues (Morris RJ et. al, 2004; Tumbar T et. al, 2004), making it a
source of adult stem cells. Hair loss (alopecia) or unwanted hairs (hirsutism) are
unpleasant side effects of aging or, more importantly, many underlying systemic

11
diseases which are important medical-sociological problems affecting the quality
of life for patients (Hadshiew IM et. al, 2004). Our increasing knowledge of skin
and HF biology help in the search for new and better treatments for management
of these diseases (Tong X and Coulombe PA, 2003; Porter RM, 2003). In
addition, skin and HFs modulate their functions with environmental changes, the
skin and its appendages are a powerful model for environmental and
developmental (Evo-Devo) research (Wu P et. al, 2004).

1.3.1 Morphology and types
There are different types of hairs on the mice based on their morphology and
location. The major ones are pelage (or trunk), tail and vibrissae hairs (Figure
1.2A). Minor hair types include these on the ears, eyelids, and perianal areas
(Sundberg JP, 2001). Hairs serve many functions including temperature
insulation, sensory, secretion, etc.

1.3.2 Structure of pelage hair follicles
Hair follicles consist of two principal tissue types: mesenchymal and epithelial.
The mesenchymal component is represented by the dermal papilla (DP) and
dermal sheath (DS). The latter is also referred to as fibrous follicular sheath. DP
is located at the bottom of the HF and has a tear-drop shape. DP is believed to
play a central regulatory role in the life of the HF, guiding its development,
growth, and cycle largely by supplying the overlying HF epithelium with
essential growth factors (Paus R, 1998).

12
Figure 1.2 Mystacial vibrissae in mice.
Comparison of pelage (left) and vibrissae (right) hair follicles. Schematic
drawing. B) Development and position of mystacial vibrissae in mice. Note the
specific position in adult mice. Panel B is adopted from Ebara S et. al., 2002

13
DP removal from the anagen HF leads to the complete cessation of the growth
activity of this follicle followed by new DP regeneration (Jahoda CA, Oliver RE,
1984a). This indicates the pivotal role of DP for the support of the HF. Anagen
DP consists of a loosely arranged population of specialized fibroblasts
interspaced by extracellular matrix (Matsuzaki T and Yoshizato K, 1998). DP
fibroblasts significantly differ from dermal fibroblasts, both by their cytological
features in vitro and by their gene expression profiles (O'Shaughnessy RF et. al.,
2004). Unlike dermal fibroblasts, DP fibroblasts show a prominent
aggregation-forming behavior in cell culture. They tend to form multilayered
clumps upon reaching a state of confluence (Jahoda CA, Oliver RE, 1984a;
Jahoda CA et. al., 1984b; Messenger AG et. al., 1986). Aggregation forming
property is retained by DP cells upon transplantation in vivo. Transplanted DP
fibroblasts remain together and regenerate a new functional DP that can even
induce a new HF when recombined with responsive epithelial cells (Horne KA et.
al., 1986). DP fibroblasts secrete a specialized set of extracellular matrix proteins
such as the SPARC-like 1 (Sparcl1), fibulin 5 (Fbln5), Osf2, chondroitin sulfate
proteoglycan 2 (Cspg2, versican), and fibronectin (Fn); signaling molecules such
as Bmp4, Igf1, Igf2, Wnt4, and Wnt5a as well as modulators of these signaling
factors (O'Shaughnessy RF et. al., 2004).

The above described anagen DP undergoes significant changes during regression
and resting stages of the hair growth cycle. The DP dramatically decreases in
volume and becomes very compact. Loss of volume is primarily accomplished
through the loss of extracellular matrix (Matsuzaki T and Yoshizato K, 1998).

14
However, migration of a subset of DP fibroblasts out into the DS can also be
accountable for the size change (Tobin DJ et. al., 2003).

The DS surrounds the HF and is connected with the DP through a structure called
the papillary stalk. DS fibroblasts are also distinct from dermal fibroblasts and
share many cytological and gene expression similarities with DP cells (Matsuzaki
T and Yoshizato K, 1998; O'Shaughnessy RF et. al., 2004, Rendl M et. al., 2005).
Additionally, at least the lower DS can regenerate a new functional DP under
certain experimental conditions and vice versa (Oliver RF, 1967; Kobayashi K
and Nishimura E, 1989; Matsuzaki T at. al., 1996). This apparent
interchangeability of DS and DP suggests a shared origin and similar biological
properties of both tissue compartments (Matsuzaki T and Yoshizato K, 1998).

The epithelial part of the HF is represented by several components. The outer
root sheath (ORS) has the shape of a hollow cylinder of epithelium with
otherwise planar polarity. The ORS is an extension of the epidermis and has a
basal layer that contacts the DS through the basal membrane around the perimeter
of the HF and suprabasal layers that contact the companion layer of the inner root
sheath (IRS). In many species beneath the HF sebaceous gland and in the area of
the arrector pili muscle attachment ORS forms noticeable anatomical protrusion -
so-called bulge (Niemann C and Watt FM, 2002b). Extensive research identified
the bulge as the stem cell niche which harbors a distinct population of
self-renewing, multipotent epithelial stem cells able to give rise to all other

15
epithelial components of the HF as well as interfollicular epidermis (Taylor G et.
al., 2000; Oshima H et. al, 2001).

Progenies of the bulge stem cells, originating from asymmetrical division, leave
the bulge and migrate along the ORS downwards before entering epithelial hair
matrix surrounding DP at the bottom of the HF. If the bulge is the repository of
the HF stem cells, then the hair matrix is the tissue factory that generates the
final products of the HF - hair shaft and IRS. Cells in the lowermost portion of
the hair matrix, below the so-called critical line of Auber (parallel to the widest
part of the DP; Auber L, 1952), proliferate rapidly. As progenies of the lower
matrix cells move upward past the line of Auber they enter the precortex
compartment of the matrix, where they start to segregate into the concentric
layers and enter the hair shaft or IRS lineage-specific differentiation program.
Distribution analysis of genotypic markers in the chimeric HFs revealed
existence of the multipotent hair matrix cells able to give rise to all hair shaft and
IRS lineages (so-called matrix stem cells, as opposed to bulge stem cells; Kopan
R, 2002). Alternatively, hair shaft and IRS lineages are derived from two distinct
progenitor matrix cells that in turn arise separately from the bulge stem cells
(Kamimura J et. al., 1997; Ghazizadeh S and Taichman LB, 2001). In the
precortex, committed cells cease proliferation and ultimately differentiate into
three distinct layers of hair fiber (medulla, cortex and cuticle) and four layers of
the IRS (cuticle, Huxley layer, Henle layer, companion layer). Cells of the hair
fiber and IRS undergo distinct differentiation programs. Cortex cells constituting
the majority of the hair shaft express distinct sets of hair keratins and keratins

16
associated proteins (KAP), and undergo trichilemmal differentiation marked by
the so-called hard keratinization (Powell B et. al., 1992; Rogers GE, 2004).
Unlike that of the hair fiber, differentiation of the IRS is associated with the
synthesis of the trichohyalin KAP resulting in so-called soft keratinization
(Rogers GE et. al., 1997; Rogers GE, 2004). Cuticle layers of both hair shaft and
IRS consist of overlapping cells pointing in opposite directions, which allows
them to interlock providing strong physical attachment between the hair shaft and
the IRS along the length of the HF up until the point just below the opening of the
sebaceous duct, where the entire IRS undergoes degradation. The companion
layer represents the forth distinct layer of the IRS and it mediates a tight
connection between the IRS and ORS (Rothnagel JA and Roop DR, 1995;
Hanakawa Y et. al., 2004).

It is interesting to compare the follicules of of different animals briefly. In other
species, animals can have differences in the composition of hair follicles. Human
and mouse hair follicles are similar. In dogs, the germinal cells are more diffusely
distributed thoughout the length of the follicle (Diaz et al., 2004). Some species
can have compound hairs in which more than one hair fiber exits a follicle. This
is not the case in humans or mice. Sometimes, more than one fiber can be seen
from the same follicular ostium. This occurs as part of the normal hair cycle
where the new actively growing hair follicle produces a new fiber which pushes
the old club hair laterally. Both the old and the new hair fibers exit from the
ostium until the old hair is lost at the exogen stage. However, in sheep, hair
follicles sharing the same osteum can branch and have multiple dermal papillae

17
and multiple hair filaments (Moore et al., 1998). Intrestingly, multiple vibrissa
follicles form in K14 noggin mouse (Plikus et al., 2004a). Bird feather follicles
appear similar to hair follicles in general structures, but these are the result of
convergent evolution. Feather follicles have a mesenchymal pulp which hair
follicles do not have, and feather stem cells are located within the follicle above
the dermal papilla (Yue et al., 2005).

1.3.3 Development based classification of pelage hair follicles
Pelage hairs are located in the skin of the trunk and are further classified into
primary (tylotrich) and secondary (non-tylotrich). Among secondary hairs awl,
auchene and zigzag types are distinguished based on their size, shape and fiber
structure (see belw; Dry E, 1926; Sundberg JP et. al., 1994b; Millar SE et. al.,
1999; Nakamura M et. al., 2001). Primary HFs start to develop at E14.5
(embryonic day 14.5), separately from secondary HFs, whose morphogenesis
does not start until E16.5 (embryonic day 16.5). In the secondary hairs, awl hairs
form in a wave before that of the zigzag hairs. While different hair types share
common signaling pathways (e.g., beta catenin, Wnt), they also depend on
different molecular pathways (Philpott MP and Paus R, 1998; Paus R and
Cotsarelis G, 1999). Primary HF morphogenesis is highly dependent on the Eda
pathway (see below; Laurikkala J et. al., 2002). Secondary HF morphogenesis
appears to depend more on the Bmp pathway (see below; Botchkarev VA et. al.,
2002a, Botchkarev VA et. al., 2002b).

18
1.3.4 Morphology based classification of pelage hair follicles
Simple hair follicle means one hair filament from one orifice. Compound hair
follicle means multiple hair filaments share one orifice. As described above (at
the end of Structures subsection), mice normally do not form compound hair
follicles, but sheeps do. The formation of compound vibrissa hair follicle in K14
noggin mice (Pllikus et al., 2004a) led us to speculate that multiple dermal
papillae can form and induce new follicles from outer root sheath. In an adult
mouse, morphological criteria are often used to identify different types of pelage
hairs. Guard hairs are straight and average 1 cm in length. They comprise only
2% of total number of pelage hairs. Awl hairs are also straight, but they are 30%
to 50% shorter than guard hairs and are much more numerous, comprising about
28% of the total number of hairs. Zigzag hairs are the most abundant type
comprising about 70% of the total pelage. They have two or more sharp bends
along the shaft. Auchene hairs are often not identified as a separate pelage hair
type due to their great variability and, in fact, similarity to the awl hairs. Looking
similar to awl hairs, auchene hairs additionally have a single bend at their distal
end. At the microscopical level pelage hairs can be distinguished based on the
number of air cell rows. Zigzag hairs have one row of air cells, guard - two and
awl and auchene hairs have two or more. While this distribution of pelage hairs is
found in most inbred mouse strains there is great variability of in groups of hairs
plucked due to site, hair cycle stage, and inbred strain (Dry E, 1926; Sundberg JP
et. al., 2001).

19
1.3.5 Structures of vibrissae hair follicles
There are several types of vibrissae in the mouse: primary (mystacial vibrissae),
secondary (supraorbital, postorbital, inter-ramal, and ulnar-carpal vibrissae), and
supernumery. Vibrissae HFs (also known as sinus/tactile HFs) are
morphologically distinct from the pelage HFs. No analogous anatomic structure
occurs in humans. Vibrissae HFs are intimately connected with the surrounding
blood sinus forming together the vibrissa follicle-sinus complex (FSC, Ebara S et.
al., 2002; Figure 1.2A). The dense collagenous capsule encloses the blood sinus
and its inner surface is lined by the mesenchymal sheath. The upper conical part
of the capsule tightly embraces the distal portion of the vibrissae follicle and is
called - the outer conical body (OCB). The vibrissal sebaceous gland is located in
the lower portion of the OCB and its duct opens into the follicle canal. Below the
sebaceous gland, expansion of the mesenchymal sheaths forms the inner conical
body (ICB) that borders the actual blood sinus. Two anatomical parts are
distinguished in the blood sinus: the upper ring sinus and the lower cavernous
sinus. The ring sinus runs around the follicle in the form of hollow ring and for
the most part has a smooth surface. In the lower third of the ring sinus does the
mesenchymal sheath form a distinct circumferential protrusion called the
ringwulst. Unlike the ring sinus, the cavernous sinus contains numerous
connective tissue trabeculae. Density, size, and convergence of trabeculae
decrease toward the follicle bulb. The follicular portion of the FSC is
anatomically similar to the pelage HFs. The dermal papilla (DP) is located at the
very bottom of the follicle. Unlike in pelage HFs, the DP in vibrissae is large and
has a distinct tip that gives it a characteristic "candle flame" shape. The vibrissal

20
DP has well defined nurturing blood vessels. The bottom of the DP is continuous
with the mesenchymal (dermal) sheath. The hair fiber and inner root sheath (IRS)
are produced by the hair matrix. The IRS ends at the level of the sebaceous gland,
while the hair fiber extends upwards through the hair canal above the skin
surface. The outer root sheath (ORS) is continuous with the epidermis. At the
level of the epidermis, the keratinocytes form a thickened region called the rete
ridge collar (RRC, Ebara S et. al., 2002; Figure 1.2A). There are vibrissae found
above the feet. These vibrissae aid in sensory perception of the environment. In
addition, the hair around the feet may remain in some mutant stocks when the rest
of the body hair is lost suggesting these are under different regulation than other
hair types.

Vibrissa serves as a somatosensory organ (Rice et al., 1986). In general, there are
nerve fibers withint the trabecules travesing the sinus. The movement of the
blood secondary to the movment of the hair induces a movement of the trabecular
area and hence gives a signal to the nerve within the trabecula. Through
trigerminal nerves and thalamus there are barrel maps in the somatosensory
contex that correspond to the arrangement of virbrissae (Durham and Woolsey,
1984). Interested cellular and molecular studies on how this this skin ectodermal
organ is connected to the brain was carried out (Kossut and Juliano, 1999;
Cybulska-Klos et al., 2004), but is the foucs of this chapter.

21
1.3.6 Functional groups of vibrissae
Vibrissae are tactile hair follicles. While no functional groups of vibrissae were
described in mice, two functionally different groups of vibrissae can be
distinguished in the mystacial pad of the closely related rats: microvibrissal and
macrovibrissal systems. Small rostral vibrissae form the microvibrissal system
with the density of follicles up to 40 times higher then that of the caudally located
macrovibrissal system, with larger HFs. The macrovibrissal system is essential
for spatial tasks, such as determining distance to an obstacle. The microvibrissal
system is important for the object recognition, but not spatial tasks (Brecht M et.
al., 1997).

1.3.7 Comparison of vibrissae among species
From species to species, vibrissae show anatomical and functional differences.
For example, vibrissae in Murinae (rodents) and Felidae (cats) exhibit significant
differences in the pattern of innervation. In rats, follicle-sinus complex
innervation is concentrated at the level of the inner conical body through the
upper half of the cavernous sinus. In cats, innervation is mostly distributed from
the rete ridge collar to the lower cavernous sinus. Predominant innervation of the
middle portion in association with the high flexibility of the vibrissae suggests
more precise tactile perception and higher acuity of vibrissae in rats than in cats.
Unlike cats, rats have poor vision and are highly dependent upon vibrissae-based
environment exploration (Ebara S et. al., 2002). Vibrissae in different species of
mammals also exhibit anatomical differences. For example, in cats, the tail of the

22
DP is extraordinary long, reaching up to the level of the ring sinus and sebaceous
gland.

In rats, the DP is limited to the hair bulb (Ebara S et. al., 2002). Unlike rodents
and cats, tammar wallaby (Macropus eugenii) from the order of Marsupialia
lacks the ring sinus and ringwhulst in its vibrissae (Marotte LR et. al., 1992).
Vibrissae of other mammals were also reported to have morphological features
different from those described above as "typical" vibrissae. Whether these
different vibrissae represent evolutionary homologues or functional convergences
remains to be studied.

1.3.8 Other types of hair follicles
Other hair types are identified based on their location on the body (Sundberg JP,
2001) and include the cilia (eyelashes), tail hairs, at least 4 types of ear hairs,
hairs around feet, external genitalia, and perianal regions.

1.3.8.1Cilia
The mucocutaneous junction of the eyelid has a single layer of long, broad hair
fibers that form a mesh network in front of the eye. These HFs are anatomically
similar to those of the trunk, but larger in diameter, and tend to remain in anagen
for prolonged periods. This is because of the longer length of the fibers, a direct
relation to the length of the hair cycle. They have a small sebaceous gland in the
normal anatomic location. Large, modified sebaceous gland are located
adjacent to these follicles; these glands are called the Meibomian glands. The

23
Meibomian gland has a single duct lined by stratified squamous epithelium that
empties through a slit-like opening onto the conjunctival surface. This gland
provides the lipid component of the fluid barrier that protects the cornea and
keeps it moist (Smith et al., 2002).

1.3.8.2 Tail hair follicles
The tail hairs are short, wide hair fibers that grow in groups of 3 in a ring-like
pattern to form concentric patterns around the tail. As these fibers emerge, the
thick epidermis of the tail is pushed into a ridge, which gives the tail the
appearance of having a series of circumferential scales along its length.

1.3.8.3 Genital and perianal hair follicles
Hair fibers around the genital openings (prepuce and vulva) and especially
around the anus are longer than those of trucal follicles. Those around the anus
produce very long fibers and have relatively large sebaceous glands. As these are
pilosebaceous unit with specialized sebaceous glands, we out them in this section.
These specialized sebaceous glands are probably somewhat analogous to the
various scent glands and perianal glands found in canids and mustellids, although
those tend to be apocrine or a combination of apocrine and holocrine glands
(Monteier-Riviere NA, 1998). In the mouse, these are large sebaceous glands that
empty via a short duct into the perianal HF.

One of the fundamental questions is the determination of regional specificity of
skin appendage types. Based on the study in the chicken, it was proposed that

24
skin Hox code may determine these specificities (Chuong, 1993). In the mouse,
the order in which the homeobox genes are transcribed may determine which hair
follicle types develop in different anatomic sites. For example, an inversion of
chromosome 15 disrupts the Hoxc cluster in hairy ears and koala mutant mice
resulting in excessive hair growth or a transition from ear hair to pelage hair
types, respectively (Sundberg JP et al., submitted).

1.3.9 Development and molecular signaling
First, we will review HF development starting from simple, flat epidermis and
finishing with the complex HF structure. Several approaches exist in the
classification of hair follicle development. Based on morphological
characteristics, stages are distinguished based on the anatomy of the developing
follicle (Hardy, 1992; Paus R et. al., 1999). These serve as important landmarks
for analyzing the pathologenesis of various spontaneous mutant and genetically
engineered mice. From a developmental point of view, creation of a HF can be
divided into three principal stages: induction, morphogenesis, and differentiation
each characterized by a distinct set of epithelial-mesenchymal interactions
associated with specific molecular signaling events (Figure 1.3).

25
Figure 1.3 Schematic representation of hair follicle morphogenesis
A) Morphological stages. Examples of signaling and structural molecules
expressed in distinct components of the developing hair follicle are listed. B)
Major developmental phases. DP - dermal papilla; SG - sebaceous gland; IRS -
inner root sheath; ORS outer root sheath. Panel A is adopted from Paus R et.al.,
1999a.

26
1.3.10 Morphological stages
1.3.10.1 Stage 0 (pregerm)
Morphologically, this corresponds to the pregerm stage (Figure 1.3A). There are
hardly any visible indications; however, some critical molecular signaling
happens in between the epithelium and underlying mesenchyme that directs both
cell types toward formation of the HF (see below). Location of the pregerms can
be visualized with the differential expression patterns of specific molecules.
Forexample, Tgfbr2 (TbetaR-II) is consistently expressed in the pregerm
keratinocytes and can be used as the useful early marker (Paus R et. al., 1999;
Paus R et. al., 1997).

1.3.10.2 Stage 1 (hair germ)
As induction proceeds, pregerm develops into the hair germ, (Figure 1.3A). This
stage is also called the hair germ stage (synonyms: germ plate, early hair germ,
follicle plug, and placode). At this stage, morphologically recognizable epidermal
thickening is seen. Basal keratinocytes assume a vertically polarized orientation.
In the underlying mesenchyme density of fibroblasts increases in the vicinity of
the hair germ and these specialized fibroblasts start to express alkaline
phosphatase (AP; Figure 1.3A; Handjiski B et. al., 1994). Interfollicular
fibroblasts remain AP negative. Again, Tgfbr2 can be used as the consistent
marker for the hair germ basal keratinocytes (Paus R et. al., 1997).

27
1.3.10.3 Stage 2
The epithelial hair germ grows downward in the form of an elongated strand of
keratinocytes (Figure 1.3A). Growth is driven by intensive proliferation within
the hair germ. The epithelial hair germ is slightly convexed at its base, which is
capped by the condensation of the mesenchymal cells of the future dermal
papillae. In Stage 2 the developing HF assumes its orientation within the skin.
The morphologically symmetrical hair germ from stage 1 most often assumes an
anterior-posterior tilt as it extends deeper into the dermis during stage 2. The
mesenchymal condensation grows under this angle with the surface of the skin
and is oriented toward the head of the developing mouse; ultimately, the
emerging hair fiber will extend caudally. Molecular markers such as Tgfbr2 and
Il1r1 (interleukin 1 receptor, type I) are expressed within the epithelial
component of the forming HF at this stage.

1.3.10.4 Stage 3 (hair peg)
The multilayered epithelial strand thickens and keratinocytes within it assume a
concentric orientation around the long, follicular axis of the hair peg. The base of
the hair peg becomes concave and encompasses a distinct ball-shaped cluster of
specialized fibroblasts called the dermal papilla (DP; synonym: follicular
papilla).
Molecular markers expressed in stage 2 can be used to label the hair peg during
stage 3 (Figure 1.3A).

28
1.3.10.5 Stage 4 (hair cone, fiber cone)
Several morphological changes occur. The epithelial hair peg continues to
elongate; its proximal end thickens and becomes bulbous. This bulb-like portion
of the hair peg partially encloses more then 50% of the DP, which itself
elongates. During stage 4 the inner root sheath (IRS) begins to form within the
epithelial portion of the hair peg immediately above the DP. It has a cone-shaped
appearance and represents the pale epithelial layer (called Henle's layer in human
anatomy) of the future mature IRS (Figure 1.3A). Molecular markers - Tgfbr2
and Il1r1 are expressed in the epithelial hair peg. DP fibroblasts are strongly AP
positive.

1.3.10.6 Stage 5 (bulbous peg)
This stage is also refrerred to: bulbous peg stage, hair canal stage, and advanced
hair cone stage). The bulge (synonym: Wulst, Cotserellis G et. al, 1990) begins to
form and is located as a group of cells that actually bulging out on one side of the
HF (Figure 1.3A). The errector pili muscle attaches in this region. Above the
bulge several enlarged cells can be seen. These are primordial cells of the future
sebaceous gland. At stage 5 these pre-sebocytes stain positive for lipid using
classical Oil Red O, Sudan black, osmium, or other lipid histochemical stains.
The bulbous part of the follicle encloses almost the entire elongated DP forming
the early hair bulb. Above the DP, epithelial cells of the precortex area continue
to differentiate. These form the complex IRS that further grows up toward the
surface of the skin. Melanocytes within the precortex area also begin to

29
differentiate and deposit melanin granules into the keratinocytes (Slominski A, et.
al, 2005).

1.3.10.7 Stage 6 (hair follicle formation)
The follicle elongates deeper into the subcutis. At the epidermal surface, the hair
canal (infundibulum) starts to form. The IRS and hair fiber, still enclosed within
IRS continue to grow towards the epidermal surface and reach the level of hair
canal. Melanin formation continues and pigment granules are deposited into the
keratinocytes of the hair fiber. Primordial sebocytes start to develop into the
sebaceous gland above the bulge region (Figure 1.3A). At this point, the
epithelial hair bulb completely encloses the now fully developed DP.

1.3.10.8 Stage 7 and stage 8
The hair fiber separates from the degrading IRS that acts as a physical barrier and
anchor for the fiber within the follicle, allowing the fiber to enter the hair canal
(Figure 1.3A). The hair fiber tip is at the level of the sebaceous gland, which by
this time is substantially enlarged and positioned above the bulge region. At stage
8, HF morphogenesis is virtually complete. The HF further grows and extends
deep into the underlying fat layer toward the panniculus carnosus (subcutaneous
muscle). The hair fiber emerges through the hair canal above the skin surface
through the follicular ostium around postpartum day 5 in most inbred mouse
strains. As such, mice are born bald. Completion of HF embryogenesis, the
so-called first hair cycle, occurs around post partum day 7.

30
1.3.11 Developmental phases and patterning
1.3.12 Inductive phase
1.3.12.1 Competence
In development, competence is defined by the ability of a tissue layer to respond
to the induction signal (Wilson R, 1999). An incompetent tissue cannot respond
even though the same induction signal is given. At the molecular level, this may
mean that signaling molecule receptors, or necessary intra-cellular signaling
molecules and transcription factors are not yet present. The HF is the end product
of a series of complex epithelial-mesenchymal interactions. Both epithelial and
mesenchymal tissues need to be competent in order to commit themselves to hair
specific developmental programs and this requires that appropriate instructive
molecular signals are provided at specific times needed during each
developmental stage. Beta-catenin is considered to be one of the molecules that
make up the competence response to interact to the dermal signal. When Catnb
(beta-catenin) was ablated in the skin using the KRT14-cre system, hair induction
was blocked, similar to what is observed in K14-Dkk1 mice (see above; Huelsken
J et. al., 2001; Figure 1.4F). By contrast, overexpression of the stabilized form of
Catnb under the KRT14 promoter in the epidermis resulted in de novo hair
formation in adult mice long after normal hair morphogenesis was completed
(Gat U et. al., 1998; Figure 1.4E). Consistently, Lef1 null mice fail to develop
pelage hairs, vibrissae, or other epithelia-mesenchymal organs, such as teeth and
mammary glands (van Genderen C et. al., 1994).

31
Figure 1.4 Examples of mutant mice with changes in pilosebaceous units
A) K14-NOG transgenic mouse (from Plikus M et. al., 2004a): Panel A1)
Appearance of control C57BL/6J (left) and K14-NOG 2.5-month-old mice. Note
the obvious hypertrichosis of the K14-NOG mouse. Panel A2, A3) Formation of
compound vibrissae hair follicles in K14-NOG mice. Panel A4, A5) Inverted skin
from the dorsal trunk region of control and K14-NOG mice. In the K14-NOG
mouse, the density of HFs is increased and the difference between primary and
secondary follicles is not obvious. Panel A6, A7) Tail skin of the K14-NOG
mouse contains an increased number of hypertrophic and dystrophic hair
follicles. Some follicles appear to have multiple dermal papillae. B) Msx2-NOG
transgenic mouse (from Kulessa H et. al., 2000): Panel B1, B2) Although hair
follicles form, the three-week-old Msx2-NOG mouse lacks external pelage hairs
and vibrissae. Panel B3-B6) During anagen Msx2-NOG hair follicles appear
wavy, and contain deposits of keratinous material beneath the epidermis. In
addition, transgenic hair follicles have loose dermal papillae and no distinct
cortex or medulla. C) K14-Cre; Bmpr1a
cl/cl
mouse (from Andl T et. al., 2004):
Panel C1, C2) Appearance of 34-days and 5-month-old K14-Cre; Bmpr1a
cl/cl

mice respectively. Panel C3, C4) Abnormal histology of dorsal skin from a
control littermate (C3) and K14-Cre; Bmpr1a
cl/cl
mutant (C4) mice at P8. Panel
C5, C6) Development of the abnormal hair follicles in 5-month-old K14-Cre;
Bmpr1a
cl/cl
mutant mice.
D) K14-Dkk1 transgenic mouse (from Andl T et. al, 2002): Panel D1) Phenotype
of F1 progeny of the K14-Dkk1 founder at P16 (mouse on the top).
Nontransgenic littermate (mouse below). Panel D2-D5) Development of all HFs

32
Figure 1.4: Continued
is arrested in the skin of high-expressing K14-Dkk1 transgenic mice (D3, D5).
Development of HFs in nontransgenic littermates (D2, D4). E) K14-∆N87
b-Cat mouse (from Gat U et. al, 1998): Panel E1) Phenotype of K14-∆N87 b-Cat
mouse ~2 months of age (mouse on the top). Nontransgenic littermate (mouse
below). Comparison of hind feet of transgenic vs. nontransgenic mouse (insert).
Panel E2-E5) Formation of epithelioid cysts and trichofolliculomas in adult
K14-Dkk1 transgenic mice.
F) K14-Cre
(neo)
; b-Cat
lox/null
mouse (from Huelsken J et. al., 2001): Panel F1, F2)
Phenotype of K14-Cre
(neo)
; b-Cat
lox/null
mice. Panel F3, F4) Loss of normal
Patched-1 expression in the skin during hair placode formation in mutant mice.
Panel F5) skin of the mutant mouse at P8; the blue line marks the area lacking
hair follicles. Panel F6, F7) Lack of hair follicles cycling in the mutant mouse
(F7).

33
Figure 1.4: Continued

34
Figure 1.4: Continued

35
Based on studies in feather induction, we suggest that the expression of beta
catenin is essential for this competence (Widelitz et al., 2000).

1.3.12.2 Dermal message
It is believed that during induction of HFs the dermal mesenchyme is the first to
initiate reciprocal molecular signaling by sending the instructive biochemical
interactive signals in the form of physicochemically optimized receptor-ligand
interactions to the already competent epidermis. The epidermis responds by
forming the hair germs (placodes). Classic transplantation experiments revealed
that the dermal mesenchyme can stimulate formation and downgrowth of hair
germs originating both from normally hair-bearing and hair-free regions of the
body (such as foot pads). In addition, mouse dermal mesenchyme is able to
induce feather buds from chick foot epidermis and scale placodes from lizard
epidermis. The regulation of the initial mesemchymal signaling is not entirely
unclear. Using the feather model, it is possible to dissociate mesenchymal cells
and overlay them with a competent epithelium. Using this model, it is shown that
the induction of placode fate results from a competitive equilibrium of
mesenchymal cell adhesion (Jiang et. al., 1999b; Jiang et. al., 2004).

1.3.12.3 Placode formation
Initially all cells in the competent epithelium have an equal ability to form
placodes. In response to the dermal message some epithelium start to elongate,
forming epidermal placodes. This may require alteration of cell adhesion and the
cytoskeleton (Jamora C et. al., 2003). One interesting and important new finding

36
is that p63, p53 homologue, plays a critical role in committing a single-layered
surface ectoderm toward a stratified epidermis during embryogenesis. The
evidence is from p63 null mice. These mice do not form placode in ectoderm. As
a result, they fail to form an apical ectodermal ridge, hair, teeth, and mammary
glands.

1.3.13 Morphogenetic phase
1.3.13.1 Epithelial-mesenchymal interactions
Induction of the HF is followed by the morphogenesis phase in which the
epithelia specified to become HF take a different path from the adjacent
inter-placode epithelia. In response to mesenchymal signaling, cells of the
epidermal placodes reply with "epithelial messages" directing underlying
mesenchyme to cluster and choose the DP fate (Sengel P, 1986; Hardy MH,
1992). Dermal papillae cells then send a second dermal message to the adjacent
hair epithelium promoting its further proliferation and HF-specific epithelial
differentiation. Unlike the first dermal message, the subsequent dermal
message does not cause morphogenetic events in the chick and lizard epithelia
and thus is specific within one class of vertebrates (Sengel P, 1976). It is
reasonable that several molecular pathways are used as mediators of
mesenchymal signaling, and several rounds of interactions are required. Some are
established, while others await discovery. Here we will highlight the cellular
events and current understanding of the major signaling pathways in the next
section.

37
1.3.13.2 Epithelial invagination
Formation of the hair germ is one of the earliest visible morphogenetic events of
hair follicle development. At the tissue level, the process of the hair germ
formation is associated with spatial rearrangement of the cells within the flat
epithelium. Changes in the cell-to-cell and cell-to-matrix attachment properties
are important prerequisites of this process. It was demonstrated that, at the
molecular level, lowering of the adhesion of the epithelial cells forming the hair
germ is achieved through downregulation of Cdh1 (cadherin 1) in these cells.
Overexpression of Cdh1 in the epidermis under the KRT14 promoter in
transgenic mice prevented required downregulation of the endogenous Cdh1,
resulting in the increased adhesiveness and formation of abnormal hair germs
(Jamora C at. al, 2003). The mechanism by which downregulation of the Cdh1
occurs, involves unusual transcriptional repression of Cdh1 by the beta-catenin
activated Lef1 transcription complexes at the level of the Cdh1 promoter. This
occurs in response to the canonical Wnt signaling within the forming hair germs.
It is also known that Wnt pathway-mediated decrease in adhesiveness of the hair
germ cells requires additional downregulation of Bmp2 and Bmp4 within hair
germs by means of the mesenchyme-derived noggin. Absence of functional
noggin protein in Nog null mice (McMahon JA et. Al, 1998) eliminated
two-thirds of hair germs normally present in mouse embryo skin at E16.5. Partial
rescue of this phenotype was achieved upon mating of the Nog null mice with
transgenic mice overexpressing Lef1 under the KRT14 promoter (Jamora C at. al,
2003; Zhou P et. al, 1995). While in the next section, we review the involvement
of molecular pathway one after another, it should be noted that morphogenetic

38
events usually require coordination of multiple pathways, such as the integration
of Wnt, Bmp, and Cdh1 (E-cadherin) signaling seen here.

1.3.13.3 Follicular formation
Early morphogenesis of the HF is associated with increased proliferation of the
hair germ and development of the conversion of mesenchymal condensations into
dermal papillae. The hair germ then grows into the dermis and also starts to wrap
around dermal condensations to form the latter dermal papilla. The epidermis
undergoes invagination to generate the outer root sheath and hair fiber. At stage
6, most follicular components are generated including inner and outer root sheath,
dermal papilla, bulge, and hair matrix which begins to generate hair fibers.

1.3.14 Differentiation phase
Starting from stage 4 (Stage 4), the inner root sheath (IRS) and hair fiber begins
to form. This requires initiation of the differentiation program within the
developing HF. This usually starts from the matrix region where keratinocytes
start to differentiate into the IRS, hair cortex (HC), and medulla. Several
mechanisms regulate the onset of differentiation. In the subsequent stages, hair
fibers have to elongate in length, and later the HFs have to go through cycling
(see section II.C).

1.3.15 Developmental patterning
In our experience, hair follicles appear to be arranged in geometric pattern. While
it is sometimes interpreted to be a random distribution, it is due more to

39
variations in observations between species than a true random pattern. In the
mouse, most investigators evaluate only vertical histologic sections in which no
patterning can be seen. When tissue is sectioned in a plane horizontal to the skin
surface, the geometrical hair follicle arrangement becomes apparent in the
laboratory mouse. Not only are hair follicles arranged in a regular pattern, the
different hair follicle types, most notably the guard hairs due to their much larger
size, are distributed in a very regular pattern. While this can be more dramatic in
species with very short hair fibers and elaborate modified skin, such as the
armadillo, mice nonetheless, have a very distinct patterning of hair follicles over
their bodies. This is further compounded by anatomic changes such as the tail and
ears, where there is a transition of hair follicle types to more specialized
structures. As with other body structures (namely segmentation in early
embryogenesis and limb formation later on) the homeobox genes play an
important role in this patterning in the skin ( Chuong, 1993; Kanzler et al., 1994;
Brancaz et al., 2004). The arrangement of hair follicles on the skin and their
orientations are set during embryonic time. Therefore the arrangement may
reflect developmental events and it is likely that certain molecular mechanisms
are responsible for setting up such a sequence.

However, the genome only gives cells a particular property, and it is the
physical-chemical interactions of the cells that lead to the pattern. In the mouse
snout, nerves do not pre-determine the pattern of vibrissa follicles (Wrenn and
Wessells, 1984). In chicken dorsal skin, Di-I labeling shows that there there is no
specific "molecular address" that put a skin appendage at a particular position. In

40
fact, in the begigging all cells are equall to begin with (Jiang et al., 1999). The
periodic patterning of skin appendages (feathers, hairs, etc.) results from reaction
diffusion. This concept was also proposed for hair by Nagorcka and Mooney
(1985) and the theoretical model developed by Meinhardt and Gierer (1974,
2000). The issues of appendage pattern formation and input of epigenetic events
were recently reviewed (Jiang et al., 2004). Recently, the Wnt receptor Frizzled 6
(Frizzled homolog 6), normally expressed in the developing and mature HFs
(Reddy ST et. al., 2004), was shown to be involved in macroscopic hair
patterning

in mice. Ablation of the Fzd6 in the Fzd6
-/-
(synonym: Fz6
-/-
) mice
results in hair patterning alterations over most

of the skin surface (Guo N et.al,
2004). While HFs are oriented toward the digits on the dorsal

surface of the feet
in WT mice, HFs form single hair whorl on each of the hind feet of the Fzd6
-/-

mice: clockwise on the right foot and counterclockwise

on the left foot. In
addition, mutant mice have aberrant hair distribution patterns on the dorsal
surface of the head, forming whorls, tufts, or ridges. In human, hair whirls
represent specific orders of placement of scalp hair follicles. Yet, the formation
of the whirl is not controlled by genetics, since homozygote twins are shown to
have one and two whirl respectively (Paine ML et. al, 2004b). Thus there is an
epigenetic component in the arrangement of skin appendage follicles (Plikus M
and Chuong CM, 2004; Jiang et al., 2004).

41
1.3.16 Molecular signaling during hair follicle development
1.3.16.1 Bone morphogenic protein (Bmp) signaling
Normal HF morphogenesis depends on balanced Bmp signaling (Botchkarev VA
et al., 1999a; Plikus M et al., 2004a). Unbalanced downregulation of Bmp2 and
Bmp4 signaling is stimulatory, while upregulation is inhibitory. Constitutive
upregulation of the Bmp pathway in Nog (noggin) null mice (Nog
tm1Am
/Nog
tm1Amc
)
affected induction of secondary HFs, and noggin from dermal papilla was
proposed to be an inducer of hair fiber production (Botchkarev VA et al., 1999a).
Morphogenesis of secondary, but not primary HFs seems to be affected by the
Bmp pathway. In vitro experiments showed stimulatory effects of noggin on HF
induction. Overexpression of chicken NOG under the KRT14 promoter in
K14-NOG mice (B6,CBA-Tg(KRT14-NOG); Plikus M et al., 2004a; Plikus M et
al., 2005) caused induction of extranumerary HFs probably through
downregulation of the Bmp pathway. Pelage hair density is increased by up to
80% in these mice. In addition, many vibrissae became compound, containing
two or three follicles instead of one (Figure 1.4A). Experimental evidence
suggests that the Bmp pathway modulates HF morphogenesis by down-regulating
the Lef1 transcription factor (Botchkarev VA et al., 1999a). Beta-catenin also
appears to be a downstream target of Bmps. Together these proteins provide
negative control of Wnt signaling (Botchkarev VA et al., 2003c). In early stages
of HF development, Bmps are involved in the induction of hair germs. Once HFs
are formed, Bmps are again involved, but this time it has a different function in
regulating differentiation in the matrix/hair fiber junction (Botchkarev VA and
Paus R, 2003a). Overexpression of the Xenopus NOG under the mouse Msx2

42
promoter in the precortex region of the Msx2-NOG mice (STOCK
Tg(Msx2-NOG) mice) disrupts normal differentiation and results in a "no hair
fiber" phenotype (Figure 1.4B). Foxn1 (the gene mutated in the nude mutation
and Hoxc13 are downregulated, while Lef1 is upregulated in the hair fiber of
Msx2-NOG transgenic mice (Kulessa H et. al., 2000).

1.3.16.2 Cut-like 1 (Cutl1) signaling
The transcriptional factor Cutl1 also promotes IRS differentiation (Ellis T et. al.,
2001). The IRS was reduced and Cutl1 downstream genes - Shh and other IRS
specific genes were down-regulated in HFs with a targeted mutation in Cutl1 in
Cutl1
Z
/Cutl1
Z
mice (Ellis T et. al., 2001).

1.3.16.3 Ectodysplasin-A (Eda) signaling
The anhydrotic ectodermal dysplasia syndrome has long been studied due to its
involvement in hair, teeth, sweat glands, etc. Mutations of ectodysplasin (EDA) in
humans result in a distinct triad of developmental abnormalities including:
anhydrosis, hypotrychosis, and hypodontia. This set of clinical features is also
known as anhydrotic ectodermal dysplasia (EDA), and it is a X-chromosome
linked syndrome, since the human EDA gene was mapped to the X chromosome
(Kere, J et. al., 1996). In mice, it was shown that only primary or tylotrich HFs
depend on Eda signaling, the mouse homolog of the human EDA gene (Vielkind
U and Hardy MH, 1996). Tabby (Ta) mutant mice lack functional ligand
ectodysplasin-A (Eda; Srivastava AK, 1997). Downless (dl) and sleek (Dl
slk
)
mice are missing ectodysplasin-A receptor (Edar; Sofaer JA, 1969). Mice with

43
crinkled (cr) mutation lack (ectodysplasin-A receptor)-associated death domain
(Edaradd; Sofaer JA, 1969). Phenotypes of tabby, downless, sleek, and crinkled
mutant mice share close resemblance to each other (Sundberg JP, 1994b). Among
many others, developmental defects of the ectodermal organs, primary, and
secondary zigzag HFs fail to develop. This results in a prominent phenotype of
naked tail and focal alopecia behind the ears (only zigzag hairs normally grow
here; Falconer DS, 1953; Sundberg JP, 1994b). While primary HFs do not form,
secondary, non-tylotrich HFs seem to develop, but based on the fiber morphology
they all resemble abnormal awl hairs. Most prominently, zigzag hairs are absent
(Green MC et. al., 1977; Sundberg JP, 1994b; Laurikkala J et al., 2002). In
addition to pelage HFs, secondary vibrissae including two supraorbital, one
postorbital, two postoral, and three interramal are affected to various degrees.
The postorbital, many postoral and interramal, and occasional supraoral vibrissae
HFs are missing (Gruneberg H, 1971; Fraser AS, Kindred, BM, 1960).
Development of HFs in homozygous and hemizygous Tabby mice begins with a
72 hours delay at E17. Also, it ends several days earlier than normal on P1-P2
(Claxton JH, 1967; Gruneberg H, 1969). The Eda/EDA (mouse/man) pathway is
homologous to Tnf/TNF pathway and its members were recently studied
extensively (Kumar A et. al., 2001; Srivastava AK et. al., 2001). The Eda
pathway provides positive control for HF induction.

1.3.16.4 Epidermal growth factor (Egf) pathway signaling
Egf signaling through the Egf receptor (Egfr) appears to be inhibitory during HF
induction (Kashiwagi M et. al., 1997). Up-regulation of Tgfa (transforming

44
growth factor alpha), whose product is a ligand for the Egfr, in the skin under the
KRT14 promoter caused the number of HFs to decrease. Similar inhibitory effect
on HF induction was achieved with Egf administration in both in vivo and in vitro
experiments (Vassar R and Fuchs E, 1991). Induction of the HF is characterized
by the mobilization of the hair germ keratinocytes within the epithelial sheath.
Cdh1 (E-cadherin) expression is downregulated and cells loose their desmosomes
and hemidesmosomes (Kaplan ED and Holbrook KA, 1994). This allows
morphogenetic rearrangements within the epidermis as HF development
progresses. Dramatic changes in the expression of signaling molecules occur
within the hair germ. Several growth factors, growth factor receptors, and
transcription factors are upregulated. In turn, mesenchymal cells of the future
dermal papilla start to express a unique set of signaling molecules. Egfr also
controls IRS specific differentiation (Hansen LA et. al., 1997). Inactivation of
either Egfr in Egfr
tm1Mag
/Egfr
tm1Mag
mice or Tgfa (transforming growth factor
alpha; Tgfa is one of the Egfr ligands) in Tgfa null mice results in premature
cornification of the IRS (Luetteke NC et. al., 1993; Hansen LA et. al., 1997).

1.3.16.5 Forkhead box N1 (Foxn1) signaling
Similar to Hoxc13, Foxn1 (formerly called the nu (nude) and Hfh11; synonym:
Whn - winged helix nude) transcription factor is expressed in hair fibers and lack
of it causes nude phenotype (Lee D et al., 1999; Figure 1.5E, 1.5F). Similar
phenotypes are results of five different allelic mouse mutations in Foxn1
(Foxn1
nu
, Foxn1
nu-Bc
, Foxn1
nu-str
, Foxn1
nu-Y
, and Foxn1
nu-StL
; Mecklenburg L et.
al., 2001). Foxn1 is believed to regulate balanced growth and differentiation by

45
promoting differentiation of the Foxn1-expression keratinocytes and proliferation
in the neighboring cells (Prowse DM et. al., 1999). Foxn1 acts as a transcriptional
regulator of multiple hair keratin genes. Krt1-3 (synonym: Ha3) hair keratin is the
most distinguished downstream target of Foxn1. Expression of Ha3 is completely
absent in pelage HFs of the nude mice. Expression of other hair keratins, such as
Ha2, Ha4, Ha5, and Ha6 are markedly downregulated (Schorpp M et. al, 2000).
Hair fibers in nude mice are twisted or locally thickened and exhibit multiple
fractures (Mecklenburg L et. al., 2001). Just like the hair fiber, differentiation of
the IRS is also controlled by Foxn1. Disruption of Foxn1 signaling in nude mice
leads to the formation of abnormal globular aggregates in the cuticle of the IRS
and cuticle of the hair shaft. In addition the hair cortex is fragmented and the
medulla is partially missing (Meier N et. al., 1999a). In turn, overexpression of
Foxn1 under the human involucrin promoter (IVL) in the IRS of the IVL-Foxn1
mice (B6;D1-Tg(IVL-Foxn1) mice) results in the wavy, curly, and truncated hair
shafts (Prowse DM et. al., 1999).

1.3.16.6 Hairless signaling
Expression of Hr (hairless) encoding a putative zinc finger

transcription factor
(Cachon-Gonzalez MB et. al, 1994) is specifically associated with the catagen
stage (Beaudoin GM et. al., 2005). Hr is expressed in part of ORS
(Krt1-14-positive), including bulge area and epithelial matrix (Krt1-14-negative).
ORS expression of Hr is maintained through late catagen into telogen and the
beginning of

the next anagen, until new hair matrix reforms (Beaudoin GM et. al.,
2005).

46
Figure 1.5 More examples of mutant mice with changes in pilosebaceous units
A) Shh null mouse (from Chiang C et al., 1999): Panel A1) Hairless but
pigmented Shh

null skin graft vs. control graft with robust hair growth. Panel A2,
A3) Inhibition of hair follicle morphogenesis in E17.5 Shh

null mouse skin (A3).
Panel A4, A5) Comparative histology of control (A4) and Shh

null skin graft. Shh

null grafts are characterized by a thickened epidermis containing keratinocyte
aggregates (arrows) at the base of the epidermis.
B) Pdgfa

null mouse (from Karlsson L et. al., 1999): Panel B1) Phenotype of P1
Pdgfa null mouse (below). Panel B2) Phenotype of P1 Pdgfa null mouse at P42
(mouse on the left). Nontransgenic littermate on the right. Panel B3) Complete
lack of anagen hair follicles in the skin of Pdgfa null mice at P25. Panel B4, B5)
Abnormal hair canals and hyperplastic sebaceous glands within Pdgfa null skin.
C) Hoxc13
neo
/Hoxc13
neo
mouse (from Godwin AR et. al., 1998): Panel C1, C2)
Phenotype of Hoxc13
neo
/Hoxc13
neo
mouse (C2). Nontransgenic littermate (C1).
Panel C3) Close-up of the mystacial pad of the Hoxc13
neo
/Hoxc13
neo
mouse
shows lack of protruding vibrissae (arrow). Arrowhead shows hair remnants
under skin surface. Panel C4) Close-up of the lower limb of Hoxc13
neo
/Hoxc13
neo

mice shows rare protruding hair. Panel C5) Hoxc13 expression in the hair based
on the beta-Gal staining of the P7 skin of Hoxc13
lacZ
heterozygote. Panel C6)
beta-Gal staining of Hoxc13
lacZ
homozygous skin counterstained with anti-K2.6
immunohistochemistry (brown).
D) Msx2
tm1 Rilm
/Msx2
tm1 Rilm
mouse (from Ma L et. al, 2003): Panel D1, D2) Skin
phenotype of the adult Msx2
tm1 Rilm
/Msx2
tm1 Rilm
mouse. Panel D3) Trunk hairs

47
Figure 1.5: Continued
from wild-type (top) and Msx2
tm1 Rilm
/Msx2
tm1 Rilm
(bottom) mice. Both club ends
are morphologically similar (arrowheads). Septation patterns are irregular in the
mutants, but there are no breakages in the middle of the shaft. Panel D4, D5)
Changes in hair filament differentiation in Msx2
tm1 Rilm
/Msx2
tm1 Rilm
mice. Medulla
patterning is affected as suggested by the irregular septations in the hair (D5).
E) Foxn1
nu
/Foxn1
nu
mouse (from Mecklenburg L et. al., 2001): Panel E1, E2)
Phenotype of the adult Foxn1
nu
/Foxn1
nu
mouse. Panel E3-E5) Dorsal skin of the
Foxn1
nu
/Foxn1
nu
mouse in early catagen (E3), telogen (E4), and early anagen
(E5). Production of a weak, twisted hair shaft results in progressive dilation of the
infundibulum encompassed by increasing amounts of cornified debris. Panel E6,
E7) Scanning electron microscopy of the 8-week-old Foxn1
nu
/Foxn1
nu
mouse
skin reveals a few twisted hair shafts emerging from the surface (E6). Hair shaft
is flattened at the bend and lacks a cuticle (E7).
F) Foxn1
tw
/Foxn1
tw
mouse (from Suzuki N et. al., 2003): Panel F1, F2)
Phenotype of the Foxn1
tw
/Foxn1
tw
mice with distinct stripe patterns. Panel F3-F8)
Histology of the skin stripe in the Foxn1
tw
/Foxn1
tw
mouse. The thick line over F3
shows the region of a black stripe. This stripe is running leftward at a speed of
1.5 mm per 30 days. Section of each position of the wave are shown in Panel
F4-F8.

48
Figure 1.5: Continued

49
Figure 1.5: Continued

50
Mutation in the Hr gene in Hr
hr
/Hr
hr
mice causes premature, highly dysregulated
catagen associated with the disruption of the hair bulb and outer root sheath into
separate cell clusters resulting in loss of epithelial contact between the upper HF
and dermal papillae (Sundberg JP et. al., 1989). The dermal papilla fails to
migrate upward and remains in the reticular dermis, where its cells undergo
apoptosis and start to express uncharacteristically high levels of adhesion
molecules. This leads to the formation of follicular cysts. In turn, upper follicular
epithelium devoid of dermal papillae transforms into utriculi (pseudocomedones;
Panteleyev AA et. al, 1999). Mutation of the vitamin D receptor (Vdr) results in
an alopecia phenotype that mimics the phenotype of Hr
hr
/Hr
hr
mice. Vdr protein
was shown to interact with the hairless protein and to cooperatively regulate
transcription (Skorija K et. al., 2005).

It was suggested that maintenance of the contact between the epithelium and DP
of the catagen and telogen HF is essential for its further regeneration during
subsequent anagen (Panteleyev AA et. al, 2000). Recent report however suggests
that maintenance of the contact between the epithelium and DP is not essential
for anagen re-entry. Attempt to rescue Hr loss-of-function phenotype by crossing
Hr
hr
/Hr
hr
mice with K14-Hr mice (STOCK Tg(KRT14-Hr) resulted in rescue of
HF ability to re-enter new anagen, but have not rescued catagen-associated hair
loss, formation of utriculi and disruption of DP / bulge contact (Beaudoin GM et.
al., 2005). It was further suggested that Hr represses expression of soluble Wnt
inhibitor (Wise) and, thus, promotes

Wnt signaling activation required for
initiation of new anagen.

51
1.3.16.7 Hox Homeobox signaling
Hox genes were shown to be important in HF growth (Awgulewitsch A, 2003).
Hoxc13 controls the expression of several genes encoding keratins and
keratin-associated proteins (Jave-Suarez LF et al., 2002; Tkatchenko AV et al.,
2001). Hoxc13 is expressed in all layers of the hair fiber and loss-of-function
mutagenesis of Hoxc13 in transgenic mice (Hoxc13
neo
/Hoxc13
neo
and
Hoxc13
lacZ
/Hoxc13
lacZ
mice) results in the absence of external hair fibers
(Godwin AR and Capecchi MR, 1998; Figure 1.5C). Overexpression of Hoxc13
also leads to fragile hair fibers (Tkatchenko AV et al., 2001). Interactions
between homeobox genes and the hairless mutation provide further insight into
patterning and hair formation (Brancaz, et al., 2004).

1.3.16.8 Msx2 homeobox signaling
The Msx2 transcription factor also plays a role in hair fiber differentiation. HFs
in Msx2 null mice (Msx2
tm1 Rilm
/Msx2
tm1 Rilm
mice) have structurally abnormal hair
shafts (Ma L et. al., 2003; Figure 1.5D) and HFs overexpressing Msx2 under the
human CMV promoter in CMV-Msx2 mice (B6;CBA-Tg(CMV-Msx2) mice) have
a thickened hyper-keratotic epidermis and a shrunken epithelial matrix region
(Jiang, T-X et. al., 1999a). This suggests it may be involved in size regulation of
epidermal thickness and the hair follicle (Wang WP et al., 1999).

1.3.16.9 Neurotrophins signaling
As HF morphogenesis progresses, proliferation is followed by the onset of hair
type lineage specification. Epithelial-derived neurotrophins prevent maturation of

52
mesenchymal condensation into the dermal papillae. This is mediated by the Ngfr
(Nerve growth factor receptor; synonym: p75NTR) expressed in the
mesenchymal condensation during stages 2 and 3 (Botchkareva NV et. al., 1999).
Ngfr-mediated signaling prevents the onset of Fgfr2 (fibroblast growth factor
receptor 2) expression associated with maturation of the dermal papillae; an event
promoted by epithelium-derived Fgf7 (fibroblast growth factor 7) Fgfr2 ligand
(Guo L et al., 1996). In agreement with this, HF development is greatly
accelerated in Ngfr null (Ngfr
tm1Jae
/Ngfr
tm1Jae
) and K14-FGFR2 transgenic mice
(which overexpresses FGFR2 under the control of KRT14 promoter (STOCK
Tg(KRT14-FGFR2); Botchkareva NV et. al., 1999).

1.3.16.10 Notch signaling
The Notch signaling pathway is involved in control of hair fiber differentiation,
postnatal homeostasis, and maintenance of the hair follicle (Vauclair S et. al.,
2005). Overexpression of Notch
∆E
(active form of Notch1) under the hair keratin
Krt1-1 promoter within the cortex in STOCK-Tg(MHKA1- Notch
∆E
) mice
disrupts differentiation in two adjacent to cortex layers: the medulla and cuticle.
This results in structural abnormalities of the hair fibers: a sheen appearance of
the pelage hair due to the loss of air spaces in the medulla, wavy pelage and
vibrissae, and brittleness of hairs resulting in visible hair loss (Lin MH et. al.,
2000). Overexpression of constitutively active form of Notch1 under the
involucrin promoter in the IRS delays IRS differentiation, and causes hair shaft
abnormalities and anagen-coupled alopecia (Uyttendaele H et. al., 2004).
Conditional loss of a Notch1 allele in the embryonic ectoderm and subsequently

53
within hair follicles in N1
flox/flox
; Msx2-Cre/+ mice resulted in a medulla defect
similar to that achieved by misexpression of Notch1 in cortex (see above). A
conditional loss of both Notch1 and Notch2 in Notch1
flox/flox
; Notch2
flox/flox
;
Msx2-Cre/+ mice or a complete loss of Notch activity in Psen1
flox/flox
; Psen2
−/−
;
Msx2-Cre/+ transgenic mice with a conditional removal of both Psen1 and Psen2
results in abnormal differentiation within HFs and conversion of the HFs into
epidermal cycts. Psen1 and Psen2 encode gamma-secretase (intracellular
protease that cuts within transmembrane domain of Notch, releasing the Notch
intracellular domain that interacts in nucleus with the transcription factor Rbpsuh,
a.k.a. RBPjk; Pan Y et. al, 2004) The outer root sheath of these mutant HFs
switch to an epidermal program of differentiation during catagen of the hair
growth cycle. Additionally, the inner root sheath cells fail to accumulate and are
replaced by large postmitotic cells (Pan Y et. al, 2004).

1.3.16.11 Platelet derived growth factor (Pdgfa) signaling
Epithelial Pdgfa signaling through its receptor Pdgfra, promotes normal
development of the dermal papillae (Kamp H et. al., 2003). HFs from Pdgfa
null mice have a small dermal papillae (Karlsson L et. al., 1999; Figure 1.5B).
Shh is thought to act as a positive regulator of Pdgfra expression within the
mesenchymal condensation. Arrested HFs in Shh (see paragraph below) null
mice fail to form Pdgfra positive mesenchymal aggregates (Karlsson L et. al.,
1999).

54
1.3.16.12 Sonic hedgehog (Shh) signaling
The Shh pathway is one of the central regulators of morphogenetic events in the
developing HF. Downregulation of the Shh pathway in various Shh null mice
(such as: Shh
tm1Amc
/Shh
tm1Amc
) arrests HF development at stage 2 preventing
progressive downgrowth of the hair germ (Figure 1.5A; Chiang C et al., 1999).
Failure of the hair germ downgrowth is partially explained by proliferative
defects in Shh null mice. This defect is in part mediated by a failure of normal
convergent Gli-mediated regulation of proliferation (Mill P at. al., 2005). Shh
regulates differentiation, maintenance of the ORS identity, and formation of the
HF stem cell compartment through upregulation of Sox9 in the ORS (Vidal VP
et. al., 2005). It is also believed that the epidermal Shh message also promotes
differentiation of the dermal papillae. In part this is mediated by Shh-induced
expression of Wnt5 within cells of the mesenchymal condensation (St-Jacques B
et. al., 1998). The canonical Lef1/beta-catenin pathway mediates Wnt5 signaling
as Lef1 expression is seen in the cells of the mesenchymal condensation starting
from stage 2 (Reddy S et. al., 2001; He X et. al., 1997).

1.3.16.13 Transforming growth factor, beta 2 (Tgfb2) signaling
Abolishment of Tgfb2 in Tgfb2 null mice resulted in significant delay and
reduction in number of induced HFs by 50%. By contrast, treatment of mouse
skin explants with Tgfb2 stimulated HF morphogenesis (Foitzik K et. al., 1999).
Interestingly, mutations in Tgfb1 and Tgfb3 genes, as well as administration of
Tgfb1 either had no effect or suppressed HF induction, suggesting complex
regulation of HF morphogenesis by the Tgf pathway (Foitzik K et. al., 1999).

55
1.3.16.14 Wnt signaling
The Wnt signaling pathway has been definitely demonstrated to be one of the
central mechanisms in promoting hair induction. Downregulation of Wnt
signaling with the Wnt pathway antagonist Dkk1 (dickkopf homolog 1) expressed
in the skin under the KRT14 (human keratin 14 promoter) blocks induction of all
hair follicle types (Andl T et. al., 2002). Expression of all other hair germ specific
markers were absent in the skin of K14-Dkk1 mice (STOCK Tg(KRT14-Dkk1);
Figure 1.4D). In addition to HFs, induction of both teeth and mammary glands in
K14-Dkk1 mice failed (Chu EY et. al, 2004). This suggests that the Wnt pathway
is among the earliest molecular events involved in HF development. Wnt
signaling through the Lef1/beta-catenin pathway (see above) plays critical role.
HFs of the Lef1 null mice fail to produce hair fibers and their development is
arrested. Regulatory sequences for Lef1 were found in the promoter regions of a
number of hair keratin genes (van Genderen C et. al., 1994; DasGupta R and
Fuchs E, 1999).

1.3.17 Comparison with vibrissa development
Unlike pelage HFs, development of the vibrissae HFs has not been investigated
in detail. The initial stages of vibrissae morphogenesis differ from that of pelage
HFs. In the literature, stage 1 of vibrissae morphogenesis is compatible with stage
1 (germ stage) for the pelage HFs. The surface epithelium grows downwards and
is surrounded by mesenchymal condensation (Van Exan RJ and Hardy MH.,
1980). However, vibrissal stage 1 is preceded by additional events not
distinguished in the pelage HFs. Unlike pelage HFs, that are initiated from the

56
flat skin, vibrissae HFs are initiated from the upward protrusion of the skin
(horizontal ridges of the "vibrissae pad", Van Exan RJ and Hardy MH., 1980). In
addition, initiation of the vibrissae seems to be intimately associated with nerve
plexus formation and mesenchymal condensation, under the presumptive follicle,
forming prior to the epithelial hair germ.

Close temporal and spatial association of the nerve plexus formation (sensory
nerve endings of the maxillary branch of the trigeminal nerve; Vincent SB, 1913;
Winkelmann, RK, 1959) beneath each vibrissae primordium further signifies the
role of innervation for the vibrissae that function as important tactile organs and
have individual projections in the barrelfield area of the somatosensory cortex
(Hoogland PV et. al, 1987; Welker E et. al., 1988). There are several types of
vibrissae in the mouse: primary (mystacial vibrissae), secondary (supraorbital,
postorbital, inter-ramal, and ulnar-carpal vibrissae), and supernumery whiskers
(Van der Loos et al., 1986; Welker and Van Der Loos, 1986; Sundberg JP, 2001).
The mystacial vibrissae are most often referred to. They can be further separated
into three major groups (Yamakado M and Yohro T, 1979; Figure 1.2B):
- nasolateral group (include horizontal rows 1 and 2);
-maxillary group, separated at E12 from the nasolateral group by the
nasolacrimal groove (include horizontal rows 3, 4 and 5);
- labial group, separated at E12 from the maxillary group by the
nasomaxillary grove;
Morphogenesis of mystacial vibrissae HFs starts as early as E12 in the "vibrissae
pad" areas located on either side of the mouse muzzle. Development proceeds in

57
the caudo-rostral direction along the five horizontal skin ridges running across the
"vibrissae pads" (Van Exan RJ and Hardy MH., 1980). Within these groups,
larger, more caudal vibrissae develop first, followed by the smaller, rostral
vibrissae (Davidson P and Hardy MH, 1952; Yamakado M and Yohro T, 1979,
Figure 1.2B). Wave-like progression of vibrissae development within the
"vibrissae pads" offers unique insight into hair development. Rostro-caudal
orientated, sections of the muzzle of developing embryos have HFs in the
consecutive stages of morphogenesis lined up next to each other (Yamakado M
and Yohro T, 1979; Van Exan RJ and Hardy MH., 1980). The position and
number of the major groups of the vibrissae HFs is strikingly constant. Each
follicle from this group can be assigned with precise coordinates (Vincent SB,
1913; Van Exan RJ and Hardy MH., 1980). By contrast, the number and position
of supernumerary vibrissae are not constant.

1.3.18 Hair cycles and molecular signaling
In adult mice, the mature hair follicle follows a cyclic, repetitive sequence of
growth activation, regression, resting, and shedding, simply known as the hair
growth cycle (Figure 1.6). Four major stages of the hair growth cycle are defined
as: anagen the period of active HF growth, catagen a transient period of
cessation of the growth and regression, telogen the period of relative inactivity,
and exogen, the point at which the old hair fiber is shed (Figure 1.6A; Dry E,
1926; Chase HB et.al, 1951; Sundberg JP, 1994a; Fuchs et al, 2001; Stenn and
Paus, 2001; Milner Y at. al, 2002).

58
Figure 1.6 Hair cycle
A) Hair growth cycle and the involvement of known signaling molecule in
transition between different stages.
B) Schematic representation of the hair follicle growth cycle stages based on
morphological criteria.
IRS - inner root sheath. Panel A is adopted from Muller-Rover S et. al., 2001.
Panel B is adopted form Botchkarev VA et. al., 2003b.

59
The hair growth cycle of pelage HFs was carefully defined and now a
classification system of the hair growth cycle stages with many substages at each
major stage is available (Muller-Rover S et. al., 2001). This classification was
defined for the C57BL/6NCrlBR strain; however, it is largely applicable to
normal mice of most if not all inbred strains and also works for other hair follicle
types (Muller-Rover S et. al., 2001). Anagen and catagen are further subdivided
into numerous stages based on a variety of morphological criteria. To date, no
distinct stages have been proposed for the telogen stage (Figure 1.6B).

1.3.19 Morphological stages of hair cycle
1.3.20 Anagen. The entire anagen stage is divided into 6 distinct stages (I-VI)
with stage III been further classified into IIIa, IIIb, and IIIc (Muller-Rover S et.
al., 2001; Figure 1.6B).

1.3.20.1 Anagen I
Launch of the proliferatory activity within the secondary hair germ and its
thickening manifests the initiation of the telogen-anagen transition and it is
identified as the first stage of the anagen.

1.3.20.2 Anagen II
The activated secondary hair germ, now termed the keratinocyte strand, further
proliferates, thickens, and elongates downwards into the dermis and hypodermal
fat layers. It partially encloses the DP that itself undergoes a noticeable

60
elargement as it approaches the dermal-subcutaneous fat border (Muller-Rover S
et. al., 2001).

1.3.20.3 Anagen IIIa
Three key of this stage are: keratinocytes form a distinct epithelial hair matrix
that encloses most of the DP; a cone-shaped IRS appears just above the DP, and
the HF crosses the dermal-hypodermal fat border. Together the hair matrix and
DP form the hair bulb. Melanin depositions become apparent in the pigmented
HFs (Chase HB et.al, 1951; Slominski A et.al, 1994).

1.3.20.4 Anagen IIIb
The new hair fiber forms. It grows downward and reaches half of the length of
the mature HF. It is fully enclosed by the IRS that by this time forms all of its
distinct layers (Henle's layer, Huxley's layer, and cuticle). During this stage the
hair bulb increases in size and is three times wider then the DP. It also contains
an abundance of melanin granules at the pole close to the panniculus muscle. The
HF extends further down to the subcutaneous fat layer.

1.3.20.5 Anagen IIIc
The hair fiber continues to grow and reaches the level just below the insertion of
the sebaceous gland. It is still fully surrounded by the IRS. At the bottom of the
HF, the hair bulb obtains its maximal sizeand reaches deep into the subcutaneous
fat layer (Chase HB et.al, 1951; Chase HB 1954).

61
1.3.20.6 Anagen IV-V
By anagen IV the hair fiber and IRS reach the hair canal. In the next stage,
anagen V, the IRS stops growing at the level of the insertion of the sebaceous
gland duct, while the hair fiber continues to grow upwards within the piliary
canal. HFs at anagen IV-V have the morphologic features of a fully developed
anagen HF and reach deep into the subcutaneous fat. It should be noted that the
subcutaenous fat layer is thickest in late stages of anagen and decreases in
thickness through catagen becoming the thinnest in telogen (Chase HB et. al,
1953; Montagna W et. al., 1951; Muller-Rover S et. al., 2001; Figure 1.6B). This
feature is often overlooked when focusing on the hair follicles and their cycles.
The increase in hypodermal fat is likely in response to the large amount of energy
needed to produce the dense hair coat.

1.3.20.7 Anagen VI
The hair fiber finally reaches the skin surface and it continues to grow through
the rest of the anagen. In general, the relative position of the hair fiber and IRS in
the HF serves as key morphological criteria in distinguishing anagen IIIc-VI
stages (Muller-Rover S et. al., 2001; Figure 1.6A). Tricholemmal plug holds the
telogen hair, which is induced by trichohyalin granules. At the level of the
sebaceous gland the inner root sheath degrades due to a variety of as yet poorly
defined enzymes released by the sebaceous gland which releases the hair fiber
and allows it to exit the hair follicle osteum (Sundberg et al., 2000).

62
1.3.21 Catagen
After the extended period of continuous growth, the HF enters the regression
phase - catagen. Eight distinct stages of catagen in mouse pelage HFs are
identified (Figure 1.6B).

1.3.21.1 Catagen I
Catagen I is morphologically indistinguishable from the preceding anagen VI.
However, during catagen I some hair matrix cells undergo early stages of
apoptosis that can be identified by immunohistochemical methods and TUNEL
(Muller-Rover S et. al., 2001; Weedon and Strutton, 1984; Lindner et al, 1997).

1.3.21.2 Catagen II
The first morphological signs of regression start to appear. Hair bulbs become
narrower and so does the DP. At the same time the papillary stalk (connecting the
DP with the dermal sheath) thickens up to three cellular layers. The number of
hair matrix keratinocytes undergoing apoptosis increases. In a pigmented HF
there is a dramatic reduction of the amount of melanin in the upper part of the
epithelial matrix (Slominski A, 1994). This results in plucked hairs lacking
pigment near the club. At the same time, there is no reduction in overall length of
the HF in comparison with the previous stages (Muller-Rover S et. al., 2001).

1.3.21.3 Catagen III
The HF shortens; the DP condenses and assumes a very distinct onion-like shape.
The hair bulb undergoes progressive involution and no longer completely

63
ensheathes the DP. In pigmented HFs, melanin disappears from the lower portion
of the follicle. Apoptosis continues on a larger scale (Lindner et al, 1997).

1.3.21.4 Catagen IV
The HF undergoes progressive shortening and the epithelial bulb further retracts
upward embossing less then 50% of the now ball-shaped DP (Muller-Rover S et.
al., 2001).

1.3.21.5 Catagen V
During subsequent catagen V the HF develops into a club hair. The depigmented
end of the developing club hair has an eosinophilic brush-like appearance. It is
surrounded by the ORS-derived secondary hair germ capsule. Formation of the
club hair and secondary hair germ capsule results in the constriction of the
developing epithelial strand between the germ capsule and compact ball-shaped
DP. The thick glassy membrane (specialized basement membrane of the HF)
appears more prominent around the constricted portion of the HF (Muller-Rover
S et. al., 2001).

1.3.21.6 Catagen VI
The brush-like proximal end of the club hair is entirely surrounded by the
secondary hair germ capsule and moves within the HF. Remnants of the IRS are
clearly seen at this period as they are sharply demarcated from the ORS. The
epithelial strand that started to form in the previous stage now elongates and is
surrounded by a glassy membrane (Muller-Rover S et. al., 2001).

64
1.3.21.7 Catagen VII
The distinguishing feature of this stage is the upward movement of the DP (Stenn
K et. al., 1998), which is coupled with the shrinkage of the proximal epithelial
component of the HF due to extensive apoptosis (Lindner et al, 1997). It is
believed that the epithelial strand pulls the DP distally, as they are tightly adhered
to each other with the help of E- and P-cadherin adhesion molecules.
Immunohistochemistry reveals that this structure contains actin filaments which
aid in this retraction. As the DP retracts, it leaves behind a trail of connective
tissue sheath (CTS) cells (Stenn KS, 2001).

1.3.21.8 Catagen VIII
HFs assume their final morphologic features which they retain until the initiation
of anagen with the next hair cycle. HFs are short and almost completely reside
within the dermis with the DP at the border of subcutis. Only the connective
tissue strand with its folded glassy membrane remains in the subcutis. Remaining
parts of the old IRS are still seen at the level of the sebaceous gland
(Muller-Rover S et. al., 2001).

1.3.22 Telogen
The telogen stage is considered a resting stage. Typically the telogen-anagen
transition is used as the starting point for the hair cycle. Right before this
transition, the HF can be easily recognized as telogenic based on the following
features. The HF is short and entirely resides within the dermis. The DP is small
and condensed. It is closely attached to the epithelial cells of the secondary hair

65
germ. Another consistent feature of telogen HFs is the absence of the inner root
sheath (IRS). Presence of the club hair (hair fiber produced during previous
anagen stage) in the HF canal above the secondary hair germ is distinct yet an
inconsistent sign of the telogen. Often club hairs are absent or are present in
multiples as they accumulate during several consecutive hair growth cycles
(Muller-Rover, 2001).

1.3.23 Exogen
Club hair shedding or exogen appears to constitute a distinct phase of the hair
growth cycle (Stenn K, 1998; Stenn K, 2005). It was demonstrated that the base
of the shed club hair differs from that of a plucked club hair. While the base of
the latter has a smooth outline rich in intact cells, the base of the former is
shrunken and more elongated with a pitted border representing deteriorated cells
(Milner Y et. al, 2002). Although shedding occurs in telogen HFs, the peak of the
exogen stage in laboratory mice is temporally coupled to anagen VI. It was
hypothesized that exogen is a two-step process. The initial phase of pro-exogen
signaling is followed by an effector stage of separation of the club hair base from
the rest of the HF. While the molecular control of the club hair shedding is
currently unknown, several pieces of evidence suggest that actual separation of
club hair during exogen is coupled with proteolytic activity (Ekholm E and
Egelrud T, 1998; Meier N et. al, 1999b; Jensen PJ et. al, 2000; Milner Y et. al,
2002).

66
1.3.24 Molecular signaling during hair cycle
1.3.24.1 Anagen-catagen transition
During this transition molecular signaling that maintained hair growth during
anagen ceases and at the same time the activity of the pro-catagen pathways
increase. It was shown that Fgf5, Tgfb1, and neurotrophins positively regulate
this catagen transition. The role of Fgf5 is clearly demonstrated in Fgf5 null mice
(both the spontaneous angora mouse mutation, Fgf5
go
, and the targeted mutation
Fgf5
tm1Mrt
) that have abnormally long hairs. In these mice, anagen is lengthened
by 3 days resulting in longer hairs (Hebert JM, 1994; Sundberg JP et. al, 1997).
The mechanism of Fgf5 action is not completely understood, however it is
possible that the short isoform of Fgf5 produced in the outer root sheath
counteracts the long form Fgf5, produced by perifollicular macrophages,
signaling on the FGFRI in the DP (Rosenquist and Martin, 1996; Suzuki S et al,
1998; Suzuki S et al, 2000). Loss of the short form Fgf5 inhibitory signaling
possibly results in late catagen entry. However, unlike Fgf5, other Fgfs, such as
Fgf8 were shown to stimulate hair growth (Kawano M et. al, 2005).

1.3.24.2 Catagen
Expression of neutrophins, such as Ntf3 (neurotrophin 3), Ntf5 (neurotrophin 5;
previously Ntf4) and Bdnf (Brain derived neurotrophic factor) are upregulated in
the proximal HF epithelium. Neutrophin receptors Ntrk2 (Neurotrophic
tyrosine kinase, receptor, type 2; synonym: TrkB) and Ntrk3 (Neurotrophic
tyrosine kinase, receptor, type 3; synonym: TrkC) start to be expressed in the DP
during the anagen-catagen transition (Botchkarev VA et. al., 1998; Botchkarev

67
VA et. al., 1999b). Modulation of the neurotrophins expression in mutant mice
signifies their role in the anagen-catagen transition. In Bdnf null mice
(Bdnf
tm1Jae
/Bdnf
tm1Jae
) or Ntf5 null mice (Ntf5
tm1Jae
/Ntf5
tm1Jae
) there was significant
catagen retardation (Botchkarev VA et. al., 1999b). Recently, a
catagen-promoting role was attributed to proNgf and p75 interactions (Peters EM
et. al., 2005). Bdnf overexpression under the alpha myosin promoter accelerated
catagen and caused a shortening of the hair length by 15% (Botchkarev VA et.
al., 1999b). Neutrophins possibly act by downregulating secretion of DP-derived
anagen promoting molecules, such as stem cell factor (Kitl, kit ligand) and
vascular endothelial growth factor A (Vegfa).

The mechanism of Tgfb1/Tgfb3 action during the catagen entry is not well
understood, however a positive role of Tgfb1 in promoting catagen has been
clearly established. Expression of Tgfb1 was shown in the ORS, while the
receptors Tgfbr2 and Tgfbr3 were shown in the ORS, IRS, and hair matrix (Paus
R et. al., 1997). Tgfb1 promotes catagen entry, while Tgfb1 downregulation in
Tgfb1 null mice (Tgfb1
tm1Doe
/Tgfb1
tm1Doe
) delays catagen development (Philpott
MP et. al., 1994a; Soma T et. al., 1998; Foitzik K et. al., 2000). It is believed that
Tgfb1 acts similar to neurotrophins by downregulating anagen
proliferation/differentiation promoting pathways. Hepatocyte growth factor
(Hgf), expressed in the DP is under Tgfb1 negative control (Shimaoka et al.,
1994; Shimaoka et al., 1995). A catagen stimulatory role was also shown for
Tgfb2 (Hibino T and Nishiyama T, 2004). Tgfb2 was demonstrated to mediate, at
least in part, the pro-catagen effect of interferon-gamma (Ito T et. al., 2005) and

68
retinoids (Foitzik K et. al., 2005). While several signaling molecules were shown
to promote catagen (see above), precise mechanism of the normal anagen-catagen
transition is not well established. Massive apoptosis is the primary driving force
of catagen. A wave of apoptosis propagates from the precortex area into the hair
matrix and then into the ORS, IRS, and hair shaft. Rapid apoptosis and tissue loss
causes collapse, creates an "apoptotic force" that leads to hair matrix shrinkage,
ORS, IRS, and hair shaft shortening (Stenn K et. al., 1998; Stenn KS and Paus R,
2001). Apoptosis in the HF during catagen is not global, but selective. HF
epithelial stem cells of the bulge, including germinative cells and transient
amplifying cells with high proliferative capabilities, and melanocyte precursor
cells are resitant to this apoptotic process due to their immune privilege and
regenerate a new HF during the next hair growth cycle (reviewed in Botchkarev
VA and Paus R, 2003a). It was shown that in human HFs status of immune
privilege is achived through very low expression of MHC Ia and suppression of
MHC II-dependent antigen presentation. Local expression of of potent
immunosuppressants (such as Tgfb1, pro-opiomelanocortin-alpha (Pomc1; aka
alpha-melanocyte stimulating hormone)) contributes to MHC I downregulation
(Paus R et. al., 2005; Ito T et. al., 2004).

The DP and dermal sheath cells do not undergo apoptosis. During catagen, DP
cells express high levels of the anti-apoptotic protein, Bcl2. They do not express
proapoptotic "death" receptors (Lindner G et. al., 1997; Matsuo K et. al., 1998;
Soma T et. al., 1998). However, the DP still undergoes significant changes. It

69
shrinks in size and the extracellular matrix is digested. Subpopulations of DP
cells migrate out into the dermal sheath (Tobin DJ et. al., 2003).

1.3.24.3 Telogen
As a resting or dormant stage follicle, telogen HFs have effective regulatory
mechanisms that allow them to maintain the inactivity status for a prolonged
period of time. The Bmp pathway is believed to play central role in telogen
maintenance (Botchkarev VA et. al., 2003c). Cells of the secondary hair germ
express Bmpr1a (Bone morphogenetic protein receptor, type 1A). It is believed
that Bmp4, which is expressed both by the secondary hair germ and neighboring
dermal papilla, acts via Bmpr1a to inhibit anagen onset (Botchkarev VA et. al.,
2001). Experimentally induced onset of the hair growth cycle can be prevented
by Bmp4 treatment. Alternatively, treatment of telogen HFs with Noggin (potent
Bmp2 and Bmp4 antagonist) stimulates anagen (Botchkarev VA et. al., 2001).
The length of telogen in wild type mice is almost twice that of the K14-NOG
mice due to the inhibitory activity of Noggin over-expressed in the skin and outer
root sheath of the HFs (Plikus M et.al, 2004a). Steroid hormones are shown to
influence the telogen status of the HFs. Estrogen was shown to exert growth
inhibitory activity on the telogen HFs via the Esr1 (Estrogen receptor 1 (alpha))
expressed in DP (Oh HS et.al, 1996; Chanda S et. al., 2000). It is believed that
expression of Bmp4 in the telogen HF is upregulated in response to estrogen
signaling (Phippard DJ et.al, 1996). In humans, androgen can act through the
androgen receptor positive dermal papilla to influence telogen/anagen length in
human scalp hair (Randall et al., 2001).

70
1.3.24.4 Telogen-anagen transition
Anagen induction is associated with the dramatic activation of multiple signaling
pathways inhibited during telogen by Bmp signaling (Botchkarev VA et.al,
2003c; Figure 1.3B). In fact, onset of anagen is associated with a marked
decrease in the expression of Bmpr1a in the secondary hair germ and increased
expression of the Bmp2/Bmp4 antagonist - Noggin in the HF (Botchkarev VA et.
al., 2001). As the overall strength of Bmp signaling weakens, Shh and Wnt
pathways are activated (Reddy S et. al., 2001). It is believed that Shh expression
in the epithelium of early anagen HFs precedes Wnt10a and Wnt10b expression
in the DP and secondary hair germ respectively (Reddy S et. al., 2001). It was
also proposed that Wnt10b in the secondary hair germ upregulates expression of
the Stat3 transcription factor (Signal transducer and activator of transcription 3;
Yamashita S et.al, 2002). Taken together, activity of these pathways stimulates
proliferation within the secondary hair germ, marking a transition of the HF into
the active growth stage, anagen. A vast array of signaling pathways operate in the
fully developed anagen HF. They control versatile proliferation, migratory,
differentiation, and secretory processes, whose coordination is essential for
proper hair formation (Botchkarev VA et. al., 2003a; Botchkarev VA et. al.,
2003b).

1.3.25 Hair follicle stem cells
Distinct populations of HF specific stem cells provide HFs with a remarkable
regenerative ability over multiple growth cycles. These stem cells reside in the
bulge of the hair follicle, located in the midportion of the HF at the level of the

71
attachment of the arector pili muscles (Cotsarelis G et. al., 1990; Oshima H et.
al., 2001). The follicular stem cells are normally slow cycling and most of the
time they remain in a quiescent state within the well-protected anatomical niche
of the bulge. Upon stimulation they are able to generate a subset of transiently
amplifying cells with high, but limited proliferative potential. Bulge stem cells
remain undifferentiated and multi-potent, as they are able to give rise to progeny
cells of various lineages within the cutaneous epithelium (Taylor G et. al, 2000
Oshima H et. al., 2001). Slow cycling is typical of stem cells. In label retention
experiments, stem cells are identified as the so-called "label-retaining cells"
(LRCs), as only they retain tritiated thymidine or bromodeoxyuridine labels after
prolonged periods of time. These labels get diluted in the fast cycling, transient
amplifying cells (Cotsarelis G et. al., 1990; Wilson C et. al., 1994; Taylor G et.
al, 2000; Braun KM et. al, 2003). Wnt signaling is implicated in maintenance of
the HF stem cells within the niche, as well as regulation of controlled exit from
the stem cell niche, and transformation into TA progenies (Lowry WE et. al.,
2005). The ability of bulge stem cells to give rise to multiple keratinocyte
lineages of the HF during the growth cycle was demonstrated in lineage tracking
studies. In the genetically engineered Krt1-15-CrePR1;R26R mice, transient
treatment with the progesterone antagonist RU496 caused conditional activation
of lacZ expression in the keratin 15-positive bulge stem cells and all of their
progeny (Morris RJ et. al., 2004). Activation of lacZ expression in the resting
telogen HFs of the adult transgenic mouse and tracing of the lacZ-labeled cells
enabled determination of the contribution of the bulge stem cells during HF
regeneration. It was shown that labeled progeny of the bulge stem cells are

72
distributed throughout all epithelial lineages of the regenerated portion of the
anagen HFs, including the hair shaft. Occasionally, lacZ-positive cells were
found within the sebaceous gland and epidermis, signifying multipotency of the
bulge stem cells and their ability to generate epithelial cell types other than hair
specific. EGFP-positive bulge stem cells isolated from the K15-EGFP mouse had
characteristically high in vitro proliferative potential, colony-forming efficiency,
and when combined with neonatal mouse dermal cells, they were able to
regenerate the entire cutaneous epithelium, including hair follicles, hair fibers,
epidermis, and sebaceous glands (Morris RJ et. al., 2004). Apart of the epithelial
stem cells, the HF harbors pluripotent neural crest stem cells (epidermal neural
crest stem cells, eNCSCs; Sieber-Blum M et. al., 2004a; Sieber-Blum M and
Grim M, 2004b). Positive for the beta-galactosidase in Wnt1-cre/R26R transgenic
mice these cells are interspaced along the length of the ORS from the bulge to the
matrix. When isolated, these cells demonstrate clonogenic ability. Clones of
eNCSCs were shown to be able to give rise to several lineages: neurons, smooth
muscle cells, rare Schwann cells, and melanocytes (Sieber-Blum M et. al.,
2004a). Presence of the melanocyte stem cells in the ORS of the HF was shown
independently in Dct-lacZ transgenic mice (Dct, dopachrome tautomerase is an
early marker of the melanocyte lineage; Nishimura EK et. al., 2002). It was
further demonstrated that hair graying is associated with the defective
self-maintenance of these melanocyte stem cells in the HF (Nishimura EK et. al.,
2005). It is likely that these melanocyte progenitor cells represent part of the
larger population of epidermal neural crest stem cells (Sieber-Blum M et. al.,
2004a). It is interesting to compare how different skin appendage follicles

73
manage their stem cells. While feather follicles evolve independently from hair
follicles, they also have feather stem cells protected in a niched region, named
follicle bulge, within the follicle. They show similar stem cell - TA (transient
amplifying) cell - differentiation homeostasis, but under different dynamics (Yue
Z. et al., 2005). This is a remarkable example of convergent evolution,
demonstrating the success of the follicle design in organs that require periodical
renewal.

1.3.26 Cycling wave of hair follicles
Hair follicles undergo constant shedding in the adult. While hairs appear to be
randomly distributed in WT mice, there are some natural growth patterns that are
revealed in mutant mice (Ma et. al., 2003; Suzuki N et. al., 2003). For example,
visualization of growth patterns can be facilitated by hair cycle dependent
changes in skin pigmentation and hair loss in mutant mice (Ma et. al., 2003;
Militzer K, 2001). In these mice, pink skin turns dark when hairs enter anagen
and turns pink again when anagen is completed. In young mice all hairs cycle
synchronously, but with increasing age the hair cycle in different regions are
asynchronous. Thus, the skin pigment pattern breaks into distinct stripes and
patches. As mice age, the stripes and patches become narrower/smaller and
eventually appear random (Militzer K, 2001). Skin pigmentation changes
progress in a wave-like fashion on the skin surface of pigmented Foxn1
tw
/Foxn1
tw

mice, allowing observation of traveling hair waves (Suzuki N et. al., 2003, Figure
1.5F). Defect in the Foxn1 gene in these mice results in the precocious
termination of the hair cycle and causes the loss of hair shafts. This is quickly

74
followed by the re-initiation of a new anagen, so that hairs cycle much faster, and
pigmentation patterns are more dynamic. In the young Foxn1
tw
/Foxn1
tw
mice, the
pigmentation oscillation takes place synchronously throughout the mouse skin.
As the mice age, the pigmentation pattern breaks into wide stripes that over time
change to narrower bands. Some mice (usually >7 months old) show narrow and
roughly evenly spaced pigmented stripes, that travel along the trunk. However,
many mice show irregular, fragmented, or very wide stripes. Msx2 null mice
(Msx2
tm1Rilm
/Msx2
tm1Rilm
) exhibit an intriguing "cyclic alopecia" phenotype (Ma et.
al., 2003, Figure 1.5D). This phenotype is due to the fact that hair fibers are
defective and are dislodged during catagen. The hairs are able to grow back.
Therefore during anagen the skin turns black and hairy, but during telogen it is
bald and pink. As the hair cycle waves advance, the alopecic regions re-enter
anagen and regain pigmentation in a progressive order. Long-term observation of
hairy and bald skin regions revealed a "cyclic alopecia" phenomenon. Hairs
within one skin domain cycle in waves but not with hairs in neighboring
domains; each domain cycles with an independent rhythm. In essence, the
"traveling stripes" of the Foxn1
tw
/Foxn1
tw
mice are the manifestation of the same
phenomenon. They represent the leading edges of hair growth domains.
Traveling stripes can also be identified in Msx2 null or classical nude mice
(Foxn1
nu
/Foxn1
nu
) by sequential photographs of pattern progression and lining
them up; however, the Foxn1
tw
/Foxn1
tw
mice bring out the waves more
prominently. Foxn1 may also be downstream to Msx2 action (Ma et al., 2003).

75
1.4 Sebaceous glands
1.4.1 Morphology of sebaceous glands
Anatomically, sebaceous glands (SG) are primarily associated with the HF and
together form the so-called "pilosebaceous unit". Each primary HF is associated
with two SGs, while secondary HFs are accompanied by a solitary SG. SGs open
into the hair canals and secrete sebum that is predominantly composed of lipids
(Paus R and Cotsarelis G, 1999; Guha U et. al., 2004). Sebaceous glands can be
simple, branched or compound alveolar in structure. Sebocytes arise from the
cuboidal lining reserve core layer to enlarge into polyhedral cells with
abundant lipid-filled vacuoles. These cells are shed into the duct where they
rupture and release their content in a holocrine mode (Monteier-Riviere, 1998).
The sebocytes lineage is distinct and separate from the rest of the epithelium
derived cells of in the skin (Merrill BJ et. al., 2001). The genetic control of
these glands has not been studied in detail. In fact, most of these glands are
routinely overlooked in phenotyping studies. The perianal and sexual accessory
glands are not found in humans and at least the preputial and clitoral glands may
be under unique regulation. Studies of allelic mutations of stearoyl-Coenzyme A
desaturase 1 (asebia, Scd1) revealed severe hypoplasia of all sebaceous and
modified sebaceous glands except preputial and clitoral glands (Sundberg JP et.
al., 2000). By contrast, null mutations in the homeobox gene Hoxd13 result in
aplasia of preputial and clitoral glands but not the other sebaceous gland types
(Johnson, KR et. al., 1998). Mutations in the ectodermal dysplasin A cascade
(Tabby, downless, and crinkled mutant mice) lack the pilosebaceous units of the
tail and directly around the ears as well as other glands.

76
1.4.2 Modified sebaceous glands
There are several kinds of modified Sebaceous glands. These include Meibomian
glands in the eyelid, cerruminous glands in the external ear cannals, preputial in
the male external genitalia, clitoral glands in the female external genitalia, and
perianal hairs and their modified sebaceous glands. Meibomian glands open
through small clefts into the conjunctiva. They produce the lipid component of
the fluid barrier that protects the surface of the cornea of the eye (Jester JV et. al.,
1988; Smith RS et. al., 2002). Like the Meibomian glands, Cerruminous glands
are very large, lobulated structures with a stratified squamous epithelial lined
duct leading into the ear canal. Preputial and clitoral glands are found at the
opening of the lower genital tracts of both male and female mice, respectively.
These are very large structures easily seen when the skin of this area is reflected.
While often misinterpreted to be sebaceous gland tumors (Nakamura Y et. al.,
2003) or teratomas (Fu L et al, 2002), these are normal structures in the mouse.
As with the other specialized glands, these are also lobulated with a stratified
squamous lined duct. Unlike the others, these accessory sexual organs undergo
atrophy and marked dilatation with age. Secondary infections result in abscess
formation which is why they are often misinterpreted to be a type of cancer. The
perianal hairs, as described under hair types, are very large structures and have
large sebaceous glands similar to those found in other hair follicle types. These
may function in scent marking as seen in other mammalian species.

77
1.4.3 Development of sebaceous glands and molecular signaling
Several signaling pathways have started to be shown to be involved in SG
development.

1.4.3.1 Bone morphogenic protein (Bmp) signaling
Bmp signaling is a known player during the cutaneous development. It is
involved in SG formation as well. Overexpression of Nog (noggin) under the rat
NSE (neuron-specific enolase) promoter in NSE-Nog mice (B6-Tg(NSE-Nog))
promoted differentiation of ectopic sebocytes in the proximal part of the follicular
ORS, away from the actual SG of the pilosebaceous unit (Guha U et. al, 2004).
The KRT14 promoter targets Nog expression differently than the NSE promoter.
Overexpression of chicken NOG under the KRT14 promoter in K14-NOG
transgenic mice (B6,CBA-Tg(KRT14-NOG) mice) led to the formation of
pilosebaceous units with hypertrophic SGs (Plikus et al, 2004a). Noggin also
suppressed formation of the eccrine sweat glands and induced ectopic
pilosebaceous units with SGs in the glabrous skin of the paws. In fact even nails
were substituted with hypertrophic skin containing SGs in K14-NOG mice.
However Meibomian glands in the eyelids of the K14-NOG mice were reduced
and many ectopic cilia formed instead. It appears that in K14-NOG mice
pilosebaceous units form at the expense of other epidermal appendages including
Meibomian glands (Plikus et al, 2004a).

78
1.4.3.2 Ectodysplasin (Eda) signaling
Among various skin appendages, SGs are also affected by the Eda pathway, the
pathway involved in ectodermal dysplasia. Tabby (Ta), downless (dl), sleek
(Dl
slk
), and crinkled (cr) mutant mice with a loss-of-function mutation in different
genes along the Eda pathway lack SGs (Gruneberg H, 1971; Vielkind U and
Hardy MH, 1996). Interestingly, both SGs associated with each primary HF are
missing, while the one SG associated with the secondary HF remain unaffected in
these mutants. Specialized meibomian glands of the eyelids are also lacking in all
of the above mentioned mutants (Gruneberg H, 1969; Sofaer JA, 1969;
Gruneberg H, 1971; Vielkind U and Hardy MH, 1996; Srivastava AK et. al.,
2001). Overexpression of the Eda-A1 (Eda isoform) either under the KRT14
promoter or under the tetracycline-responsive promoter elements (TRE) exerts a
hypertrophic effect on the SGs (Cui CY et. al., 2003; Mustonen T et. al., 2003).

1.4.3.3 Hedgehog signaling
The effect of Bmps inhibition on sebocyte differentiation may be mediated by the
activation of the downstream Shh pathway (Allen M et. al., 2003) that was shown
to be essential for SG differentiation as well. Shh signaling increases both size
and number of SGs and causes formation of ectopic sebocytes (Allen M et. al.,
2003). Experimental inhibition of hedgehog signaling by overexpression of the
dominant negative Gli2 (Gli2∆C4) under the bovine KRT5 promoter selectively
suppressed sebocyte development. By contrast, activation of hedgehog signaling
in transgenic mice overexpressing a constitutively activated

form of a hedgehog
pathway signal transducer smoothened homolog (Smo; activated form -

79
M2Smo) under the KRT5 promoter greatly increased the number and size of SGs
and induced ectopic SGs not associated with the HFs in the footpad epidermis.
These SGs secreted sebum directly onto the surface of the skin (Allen M et. al.,
2003).

1.4.3.4 Myc (myelocytomatosis oncogene, c-myc) signaling
C-Myc is a known downstream target of Wnt signaling (Honeycutt KA and Roop
DR., 2004). Overexpression of Myc under the KRT14 promoter causes
overproliferation in the epidermis and epithelium derived structures and
ultimately depletion of the epidermal stem cells (Honeycutt KA and Roop DR.,
2004). As cells exit the stem cell state, and in mutant mice they preferentially
differentiate into sebocytes and "inner follicular epithelium". However, the
possibility exists for the observed effect of c-Myc to be secondary. c-Myc
overexpression reduces the amount of Itgb1 (integrin beta 1) expression and
decreases cell migration. It can not be excluded that cells unable to effectively
migrate from the bulge remain in close vicinity of the SG and are influenced by
the SG-specific morphogenetic factors (Waikel RL et. al., 2001; Arnold I et. al.,
2001).

1.4.3.5 Peroxisome proliferator activated receptor gamma (Pparg) signaling
Transcription factor Pparg appears to be essential for the differentiation of the
SGs. In Pparg null/WT chimeric mice SG forms exclusively from the WT cells
(Rosen ED et. al., 1999). Consistent with this observation, Pparg ligands promote

80
sebocyte differentiation in vitro (Rosen ED et. al., 1999; Rosenfield RL et. al.,
2000).

1.4.3.6 Wnt signaling
Wnt signaling was shown to be essential in sebocyte-specific fate determination
of the epidermal cells. Inhibition of the Wnt response in mice overexpressing a
dominant negative variant of the Lef1 under the KRT14 promoter in K14- ∆NLef1
mice (B6;CBA-Tg(KRT14-∆NLef1)) blocked HF maturation and promoted
sebocyte differentiation. K14- ∆NLef1 mice developed sebaceous adenomas
originating from the upper HF region and sebomas with the elements of
sebaceous differentiation (Niemann C et. al., 2002a; Niemann C et. al., 2003).

1.5 Sweat glands
1.5.1 Morphology of sweat glands
In mice eccrine glands are present only in the foot pads. Secretion of these glands
appears to increase tactile sensation and traction on the foot pads. Due to its
limited distribution and number, the eccrine glands thermoregulatory role in
mice is insignificant (Taylor JK et. al, in press). Eccrine (merocrine) glands
consist of a highly coiled secretory portion and ductal portion. The upper, distal
segment of the duct is referred as the acrosyringium. It penetrates the epidermis
and opens on its surface as the glands aperture (Sato K et. al., 1989). One type of
cells, equivalent to clear cells is lining secretory acini of the mouse eccrine
glands (Quick DC et. al., 1984). Myoepithelial cells surround the secretory
portion of the glands on the periphery. Developing eccrine glands become first

81
morphologically apparent at E16.5 in the form of placode-like cellular
accumulations immediately beneath the epidermis. By E18.5 these placodes
invaginate into the dermis and form the ductal part of the gland. Eccrine glands
develop acrosyringium by P6-7 and by P21 they reach the morphology of the
adult, a fully developed sweat gland (Taylor JK et. al, in press). There are also
apocrine glands which are simple saccular or tubular glands frequently opened to
the hair follicle, and produce more viscous secretions (Monteier-Riviere NA,
1998). In domestic animals, the apocrine glands are widely distributed over the
skin, in contrast to the human in which they are mainly in the axillary, pubic, and
peri-anal region.

1.5.2 Development of sweat glands and molecular signaling
Not much is known. Mice with disrupted Eda signaling (crinkled, downless,
sleek, and tabby) completely lack eccrine glands (Figure 1.7; Srivastava AK et.
al, 1997). This resembles the absence of sweat glands in human with some types
of anhidrotic ectodermal dysplasia (Blecher SR, 1986). In K14-NOG mice the
majority of the eccrine glands are developmentally substituted with pilosebaceous
units (Plikus M et. al, 2004a).

1.6 Nails
1.6.1 Morphology of nails
Mammals form cornified skin appendages on the tips of the digits. In mice, like
in many other mammals, they are known as claws, or nails. However several

82
Figure 1.7 Sweat glands
A) K14-EDA-A1 mouse (from Mustonen T et. al., 2003): Panel A1 - A3)
Increased sweat production as visualized by iodine-starch staining reflects
stimulated formation of eccrine glands in K14-EDA-A1 mice (A2) vs. wild type
(A1). Tabby mice have almost no staining (A3).

83
morphological modifications exist. They are known as nails in humans or hooves
in horses (Hamrik MW, 2003). Mouse nails have longitudinal and lateral
curvature. The nail plate extends distally well beyond the tip of the digit.
Proximally it is demarcated from the rest of the digit by the proximal nail fold.
Proximal nail fold extends laterally and delineates the nail plate from the sides.
At the edge of the proximal fold is the eponychium. Ventrally the nail plate is the
wide zone of hyponychium that further continues into the foot pad. At the
proximal base of the nail plate is the zone of actively proliferating epithelial cells,
known as nail matrix. As the nail matrix cells exit the proliferation state they
move distally and terminally differentiate. The mature nail plate consists of
superficial and deep layers of hard keratin. (Figure 1.8, Figure 1.10)

1.6.2 Development and molecular signaling
The first morphological sign of the developing nail is the primary nail field in
form of thickened epithelium on the dorsal side at the tip of the digits at E14. At
E15 the proximal transverse groove further delineates this nail placode from the
rest of digital epidermis dorsally and laterally. As the transverse groove folds
deepen at E16, the basal epidermal cells at the bottom of the fold develop into the
germinal matrix of the nail. The proliferating germinal matrix will later become
the primary source of the nail cells that will differentiate and keratinize as they
move distally along the nail bed. Upon or slightly after birth the nail plate reaches
the end of the digit. At that time nail is morphologically similar to that of the
adult (Figure 1.8).

84
Figure 1.8 Nail development
Stages of the mouse nail development. Scanning EM and sagittal sections.
Adopted from Bereiter-Hahn J et. al., 1984.

85
1.6.2.1 Bone morphogenic protein (Bmp) signaling
Nail development is influenced by fluctuations in Bmp signaling. Inhibition of
Bmp2/Bmp4 signaling in K14-NOG mice causes nail agenesis (Plikus M et al.,
2004a). The nail induction fails and the nail fields undergo alternative epidermal
differentiation. K14-NOG mice with low copy number of the transgenic construct
develop aberrant nails instead. Their nails are split into several nail plates, and
show abnormal differentiation with a loss-of-function. An excess of noggin
causes excessive proliferation in the nail matrix and delays nail-specific
differentiation (Figure 1.9B). A nail phenotype was also reported for the
Msx2-NOG transgenic mouse. It is interesting to note that NOG expressed under
the Msx2 promoter causes dorsalization of the digit tips and often causes
formation of the supernumerary ventral nails with mirror image symmetry
(Kulessa H et. al., 2000; Wang CK et al., 2004; Figure 1.9C).

1.6.2.2 Eda signaling
In mice overexpressing Eda-A1 under the KRT14 promoter nails grew longer than
normal (Mustonen T et. al., 2003).

1.6.2.3 Foxn1 signaling
Nude mice bearing a mutation in the Foxn1 have largely downregulated
expression of several hair keratin genes within the HF, resulting in a prominent
hair fiber and IRS dysplasia. In addition nude mice have nail defects (Flanagan
SP, 1966).

86
Figure 1.9 Mutant mice with changes in nails
A) Adult nails of Hoxc13 null mice are malformed. They have a flatter
appearance, and are often twisted (from Godwin AR and Capecchi MR, 1998).
B) K14-NOG transgenic mouse (from Plikus M et. al., 2004a): Panel B1, B2)
Aberrant nails in K14-NOG mice with a low copy number of the transgenic
construct. Nails are split into several nail plates and show abnormal
differentiation with a loss-of-function. Panel B3) Agenesis of nails in K14-NOG
mice with a high copy number of the transgenic construct. Nail induction fails
and the nail fields undergo alternative epidermal differentiation.
C) Nail phenotype of the Msx2-NOG transgenic mouse. Expression of NOG
under the Msx2 promoter causes dorsalization of the digit tips (Panel C2) and
often causes formation of the supernumerary ventral nails with mirror image
symmetry (Panel C3; from Wang CK et al., 2004).

87
Figure 1.9: Continued

88
Onychocytes within the nail matrix of nude mice undergo abnormal
differentiation. Suprabasal onychocytes do not express Krt2-1 (keratin 1), but
instead contain ectopic filaggrin-containing keratohyalin granules. Altered
onychocyte differentiation causes them to cornify and separate from the
hyponychium. As a result nude mice nails are short and unusually thin
(Mecklenburg L, 2004). Recently nail dystrophy in humans was associated with a
heterozygous mutation in

the FOXN1 gene (Auricchio L et. al, 2005).

1.6.2.4 Hairless and vitamin D receptor
Many allelic mutations in the hairless gene (Hr; hairless and rhino mutant mice),
vitamin D receptor targeted mutation (Vdr
tm1.Gcm
), and ornithine decarboxylase (a
member of putricine cycle) transgenic mice (C57BL/6-TgN(K6ODCtr)55Tgo) show
similar phenotypes in the nail (Zinser et al., 2002; Panteleyev et al., 2000). They show
long, curly nails (onychocryptosis) due to separation of the nail plate from the
nail bed. This pathology is because of the extension of the cornified epidermis
under the nail plate. It is best demonstrated by the extension of the granular layer
under the nail plate as shown by antibodies to filaggrin.

1.6.2.5 Hoxc13 signaling
During the nail development Hoxc13 shows localized expression in the forming
nail and is thought to control the expression of various keratin genes. Adult nails
of Hoxc13 null mice are malformed. They have a flatter appearance, and are often
twisted (Godwin AR and Capecchi MR, 1998; Figure 1.9A).

89
1.6.2.6 Msx2 signaling
Msx2 appears to be important for normal growth of the nail. While nails in the
Msx2 null mice form they are enlarged and more curved (Satokata I et. al., 2000).

1.6.2.7 Notch signaling
Expression of ectopic Notch1 in postmitotic cells within the keratogenous zone in
mice, results in elongation of the nails. This is achieved non-autonomously and is
indirectly mediated by Notch1 expansion in the proliferating nail matrix (Lin MH
and Kopan R, 2003).

1.7 Volar pads
1.7.1 Morphology and types of volar pads
Mouse volar skin forms various protrusions that constitute basic dermatoglyphic
patterns. Many large and small protrusions (pads) can be identified on the surface
of the palm and sole of the mouse. These pads are classified into four types
(based on Tsugane M and Yasuda M, 1995; Figure 1.10C, 1.10D):

- digital pads. Digital pads are single, large pads at the tip of each digit. They
contain many eccrine sweat glands. Dermal ridges surround the sweat glands and
collectively form a finger print-like pattern, often in the shape of a honeycomb.
However this pattern of dermal ridges is obscured by a thick layer of keratinized
epidermis.

90
Figure 1.10 Development of the mouse volar (foot) pads
A) Scanning EM of the developing foot pads.
B) Proliferation (shown by arrows) in developing foot pads.
C, D) Dermatoglyphic patterns of the pads on mouse volar skin.
E, F) Cross-section through the mouse digit and volar pads.
d - digital pads; c- carpal pads; ca - caterpillar pads; co cobblestone pads;
h-hair follilcle; id -- interdigital pads; sd sebaceous duct; sg sebaceous
gland; t- tarsal pads. Panel A, B from Bereiter-Hahn J et. al., 1984). Panel C-F
from Tsugane M and Yasuda M, 1995.

91
Figure 1.10: Continued

92
Figure 1.10: Continued

93
- interdigital pads and carpal (tarsal) pads. There are three interdigital and two
carpal pads on the palm. Carpal pads are about three times bigger than interdigital
pads. On the sole, there are four interdigital and two, similarly sized tarsal pads.
A deep, visor-like distal fold accompanies every interdigital pad. Each pad has
multiple sweat glands, dermal ridges, and tongue-like dermal projections.

- caterpillar pads. Caterpillar pads, named so after their look, form a series of
elongated protrusions parallel to the long axis of the digit. Caterpillar pads are
present on all but the first forelimb finger. Otherwise, the number of the
caterpillar pads ranges from four to nine, been is constant in palm fingers and
variable in the sole fingers. Similar to other pads, caterpillar pads contain sweat
glands, honeycomb-shaped dermal ridges, and tongue-like dermal projections.

- cobblestone pads. Cobblestone pads are the smallest pads interspaced around
interdigital and carpal (tarsal) pads. It is notable that proximal caterpillar pads
tend to break up and gradually transform into the cobblestone pads. Each small
cobblestone pad has at most two sweat gland openings.

1.7.2 Development of volar pads and molecular signaling
Condensation of the mesenchymal cells beneath the pad epidermis becomes
apparent at E13 and serves as the first indication of the developing footpads.
Interdigital and tarsal pads of the hind limbs start to form first. In E14 hind limb
pads, mesenchymal condensations become visible as slight protrusions of the
volar skin. Condensation of the mesenchymal pad cells does not appear to be

94
driven by proliferation up until E15.5, when high, and localized proliferation
activity is seen directly beneath the pad epidermis. High proliferation rates in the
mesenchyme stop by E16.5. At the same time the footpad epidermis starts to
thicken and sweat ducts start to form. At E18 dermal ridges of the footpads begin
to form fine tongue-like projections that are getting populated by peripheral nerve
endings. These processes continue into the postnatal period (Mori N et. al., 2000;
Figure 1.10A, 1.10B).

1.7.2.1 Bmp signaling
Proper Bmp signaling appears to be important for proper footpad development.
Downregulation of the Bmp2/Bmp4 activity in the volar skin of the K14-NOG
mice causes hypoplasia of the footpads, yet does not change their total number.
Noggin suppresses the localized

mesenchymal cell proliferation that is otherwise
positively

controlled by Bmps (Plikus M et.al, 2004a). Noggin also suppresses
sweat glands

and causes alternative formation of HFs and sebaceous glands in the
volar skin (Figure 1.11B).

1.7.2.2 Engrailed 1 (En-1) signaling
En1 is a homeodomain-containing transcription factor. Normally it is expressed
in embryonic ventral limb ectoderm (Loomis CA et. al., 1996). Mice deficient for
functional En1 show dorsalization of the ventral paw as well as a patterning
defect along the proximal-distal axis (Loomis CA et. al., 1998). Specifically,
digital pads at the tips of the fingers are replaced with the circumferential,
cylindrical, or doubled nail (Figure 1.11C).

95
Figure 1.11 Mutant mice with abnormal volar pads
A) Wnt7a null mouse (from Parr BA and McMahon AP, 1995): Panel A1, A2)
Dorsal and ventral sides of the paw of the control mice. Panel A3, A4) Formation
of ectopic pads in the dorsal skin of the limb of Wnt7a null mice. However,
unlike ventral pads (A3), dorsal pads are pigmented (A4). B) K14-NOG
transgenic mouse (from Plikus M et. al., 2004a): Panel B1, B3) Volar pads of
control mice. Panel B2, B4) Hypoplasia of the volar pads in K14-NOG mice with
a high copy number of the transgenic construct. Panel B5, B6) Suppression of
sweat glands

and alternative formation of HFs and sebaceous glands in the volar
skin of K14-NOG mice with a high copy number of the transgenic construct (B6).
C) En1 null mice (from Loomis CA et. al., 1996): Panel C1, C3) Whole mount
view of the ventral side (C1) and cross-section (C3) through the limb of adult
control mice. Panel C2, C4) Dorsalization of the ventral paw as well as a
patterning defect along the proximal-distal axis in En1 null mice. Digital pads are
replaced with the circumferential, cylindrical, or doubled nail (C4).
D) Severe volar pad aplasia in Hoxd13
spdh
/ Hoxd13
spdh
mice (D2) in comparison
with control (D1; from Hamrick MW, 2003).

96
Figure 1.11: Continued

97
Interdigital pads form in the mutant animals, but they have few if any sweat
glands, become pigmented, and even, partially differentiate into a nail-like
structure, forming unusual pad/nail hybrid structures. Additionally, ectopic hairs
form in the ventral skin of the En1 null mice (Loomis CA et. al., 1998).

1.7.2.3 Hox signaling
Expression of the posterior Hox genes, such as Hoxd13 and Hoxc13 coincides
with footpad formation. Hoxd13 null mice (Hoxd13
spdh
/ Hoxd13
spdh
) show severe
footpad aplasia (Hamrick MW, 2003). It is believed that Hox genes control
normal volar pad growth through the regulation of mesenchymal proliferation
(Figure 1.11D). Administration of retinoic acid (RA) to mice at E12.5 causes
aplasia of some footpads and hypoplasia of those that form. Reduced
proliferation is thought to be the underlining mechanism of the foot pad
phenotype in RA-treated animals (Mori N et. al., 2000). It is believed that RA
action is partially mediated by Hox genes, since RA is known to bind to the
enhancer regions of the Hox genes and modulate their expression (Hamrick MW,
2003).

1.7.2.4 Wnt signaling
In Wnt7a null mice mesenchymal cells undergo dorsal-to-ventral fate
transformations (Parr BA and McMahon AP, 1995). As a result, ectopic footpads
form in the dorsal skin of the limb. However unlike ventral footpads, dorsal
footpads are pigmented in Wnt7a null mice? (Figure 1.11A).

98
1.8 Mammary glands
1.8.1 Morphology of mammary glands
Mammary glands are often referred to as modified sweat glands and thought to
derive from apocrine-like glands of synapsids (Oftedal OT, 2002). In specied
with multiple mammary glands, the glands develop the along mammary lines,
also called milk lines (Veltmaat JM et. al., 2003). In mouse, there are five pairs of
mammary glands. At E11.5 five placodes or mammary anlagen form along each
milk line as a slightly protruding lens-like thickening of ectoderm.

The post-pubertal mammary gland is composed of a branching ductal system
ending in specialized lobules. The entire mammary tree is lined by an inner layer
of luminal epithelial cells surrounded by an outer layer of myoepithelial cells. It
appears that these two layers originate from a common ectodermal stem cell
located in the ductal luminal compartment (Chepko G and Smith GH, 1997;
Ferguson DJ, 1985; Kordon EC and Smith GH, 1998; Pechoux C et. al., 1999;
Stingl J et. al., 1998). It is believed that cap cells (precursors of myoepithelial
cells (Humphreys RC et al., 1996) of the terminal end bud also represent progeny
of the ductal stem cells (Kordon EC et. al., 1995). The concept that ductal stem
cells are the progenitor cells of the entire mammary gland is supported by
autoradiographic studies showing that these cells cycle at a lower pace than any
other population of mammary cells (Smith GH and Medina D, 1988; Welm BE
et. al., 2002; Zeps N et. al., 1996). Moreover, they can regenerate a complete
mammary tree when transplanted into mammary fat pads (Welm BE et. al.,
2002).

99
1.8.2 Prenatal development and postnatal changes in mammary glands
1.8.2.1 Prenatal development
By E13 placodes develop into buds that invaginate into the mesenchyme (Figure
1.12). Early bud growth is predominantly driven by cell migration, rather than
proliferation (reviewed in Veltmaat JM et. al., 2003). The mesenchyme
surrounding the epithelial buds undergoes change and forms a dense
condensation, referred to as mammary mesenchyme. The latter is distinct from
the surrounding dermis and is thought to send instructive morphogenetic signals
to the epithelial placodes (Kratochwil K et. al., 1969). Experimentally the
mammary mesenchyme can induce new mammary glands in the epithelium
otherwise not involved in mammogenesis. In E15.5 female embryos proliferation
rises at the distal tip of each mammary bud causing the outgrowth of a primary
epithelial sprout into the presumptive fat pad (Figure 1.12). The lumen of the
primary milk duct forms on the proximal end of the primary sprout and opens to
the outside. Starting from E16 the primary sprout starts branching
morphogenesis. At about the same time the epidermis surrounding opening of the
primary milk duct invaginates and forms a circular nipple sheath, that later
matures into the nipple. The mammary mesenchyme
controls nipple formation, and nipples are absent in adult male mice, whose
mammary mesenchyme degenerates early (reviewed in Veltmaat JM et. al.,
2003). By E18.5 the mammary gland has a well-formed small ductal tree with
about 10-20 individual branches surrounded by fat pad. The fat pad is the
connective tissue distinct from the mammary mesenchyme and in adult females it
forms the mammary gland stroma (Figure 1.12).

100
Figure 1.12 Stages of the mouse mammary gland development
Epidermis, yellow; dermal mesenchyme, blue; fat pad, green. Embryonic days
are shown. Dm- dermal mesenchyme; fpp fat pad precursor; mm mammary
mesenchyme; ns nipple sheath. From Veltmaat JM et.al., 2003.

101
Fat pad precursor tissue is essential for mammary epithelium growth and
branching morphogenesis (reviewed in Veltmaat JM et. al., 2003).

1.8.2.2 Postnatal changes
Development of the mammary glands in females does not stop upon completion
of embryogenesis, but continues throughout puberty, and pregnancy (Daniel et
al., 1987; Sakakura T et. al., 1987). Until puberty, growth of the mammary gland
is hormone-independent. Hormone-dependent growth starts upon puberty,
during the 6-8
th
week after birth. Rapid ductal elongation and side branching
occur. Pregnancy stimulates the development of lobular-alveolar structures for
lactation. After weaning, mammary gland involution starts, and is associated with
the loss of alveolar structures (Strange R et al., 1992). Terminal differentiation
markers throughout the development of the mouse mammary gland undergo
dynamic changes. The expression of alpha-smooth muscle actin (SMA) and
cytokeratins Krt2-5 (human synonym - CK5), Krt2-8 (human synonym CK8)
and Krt1-14 (human synonym CK14) are influenced by the development stage
of the mammary gland. Expression of Krt2-5 and SMA is restricted to the basal
cells. Cytokeratin Krt1-14 is consistently expressed by the mammary basal cells,
and is also detected in the scattered luminal cells from E13.5 through puberty.
Labeling for Krt2-8 in luminal cells is heterogeneous at all times. Heterogeneous
expression patterns in luminal cells suggest this layer has cells with a variety of
biological functions. Cytokeratins 1, 6, 10, 13, and 15, filaggrin, involucrin, and
loricrin are not detected at any stage of mammary gland development (Mikaelian
I et. al., submitted).

102
1.8.3 Molecular signaling during mammary glands development
During their development, mammary glands utilize similar signaling pathways as
do other skin appendages. However, a conclusive role in mammogenesis was
demonstrated only for few signaling pathways.

1.8.3.1 Fgf signaling
Fgf pathway activity is essential for the early mammogenesis (Dillon C et. al.,
2004). Members of the Fgf signaling pathway are expressed in the developing
mammary glands with Fgfr2(IIIb) seen in the mammary placode as early as E11.
One inguinal and three thoracic and placodes never form in both Fgf10 null and
Fgfr2(IIIb) null mice (Figure 1.13A, 1.13B). First inguinal mammary anlagen
forms. However in Fgfr2(IIIb) null mice it reaches and stops at the bud stage and
in Fgf10 null mice it progresses further but fails to branch (Mailleux AA et. al,
2002). Based on the Fgf10 expression in the fat pad cells, Fgf10 role in
adipocyte-specific differentiation and branching defect of mammmogenesis in
Fgf10 null mice, Fgf10 was proposed as the key mediator of fat pad signaling to
the primary sprout (reviewed in Veltmaat JM et. al., 2003). Fgf signaling during
early mammogenesis is in part mediated by downstream activation of Tbx3
expression (Eblaghie MC et. al., 2004).

103
Figure 1.13 Mutant mice with abnormalities in mammary glands
A) Fgfr2(IIIb) null mouse (from Mailleux AA et. al, 2002) : Panel A1-A4)
Normal Lef1 expression in the area corresponding to mammary bud 4 (A1, A3).
Lef1 expression is not present in the corresponding area in the Fgfr2(IIIb) null
embryo (A2, A4). Panel A5, A6) High number of apoptotic cells labeled in the
epithelium of the E12.5 Fgfr2(IIIb) null mammary bud (A6) detected by TUNEL
method. Normal mammary bud at this stage showing a complete absence of
apoptotic cells (A5) stained by the TUNEL method.
B) Fgf10 null mouse (from Mailleux AA et. al, 2002): Panel B1-B4) Absence of
all but mammary bud 4 in the Fgf10 null E12.5 embryo, as visualized by Lef1
expression (B2 vs. B1). Developing mammary bud 4 in Fgf10 null embryos show
normal morphology (B4 vs. B3). Panel B5-B8) Normal ramification of the
epithelium of mammary gland 4 from the Fgf10 null mouse in comparison with
control mammary gland (B7, B8 vs. B5, B6).
C) Pthrp null mouse (from Wysolmerski, JJ et. al, 1998): Panel C1, C2) Primary
epithelial duct is seen arising from the epidermis and extending below the dermis
in mammary glands from control E18 embryos (C1). In Pthrp null E18 embryos
the epithelial duct does not extend out of the upper dermis and becomes
surrounded by an abnormally dense condensation of fibroconnective tissue
(arrow, C2). Panel C3, C4) Whole-mount analysis of control 4-month-old
mammary gland 4, shows a fully branched epithelial duct system surrounding the
central lymph node (C3). At the same time, the mammary gland from the Pthrp
null mouse is devoid of epithelial structures (C4).

104
Figure 1.13: Continued
D) K5-rtTA, tetO-Dkk1, TOPGAL (Dkk1-expressing/TOPGAL) mouse (from Chu
EY et. al, 2004): Panel D1, D2) Strong staining for beta-galactosidase reveals
TOPGAL Wnt reporter expression at sites of mammary placode development in
E11.5 control embryos (D1). TOPGAL activity is severely reduced in mice
overexperssing Dkk1 under the control of the tetracycline/doxycycline-responsive
promoter (D2). Panel D3, D4) Mammary placodes are present in E11.5 control
embryos and are absent in Dkk1-expressing embryos as revealed by scanning
electron microscopy (D3 vs. D4).
E) Tbx3
tm1Pa
/Tbx3
tm1Pa
mouse (from Davenport TG et. al., 2003): Panel E1, E2)
Absence of mammary buds in E11.5 Tbx3
tm1Pa
/Tbx3
tm1Pa
embryos as evidenced
by Wnt10b expression (E2 vs. E1). Panel E3, E4) Absence of mammary buds on
sagittal sections of E12.5 Tbx3
tm1Pa
/Tbx3
tm1Pa
embryos (E4 vs. E3).
F) K14-EDA-A1 mouse (from Mustonen T et.al., 2003): Panel F1-F3) Unlike
high and tapering nipples of control female mice (F1), nipples of K14-EDA-A1
transgenic females are round. Additional rudimentary nipples are formed (F2).
Some of them can be associated with a small mammary gland fat pad (F3).

105
Figure 1.13: Continued

106
1.8.3.2 Eda signaling
Ectodysplasin-A (Eda) signaling was shown to be important in the early
patterning of the mammary placodes along the milk line. Unbalanced
overexpression of Eda-A1 (Eda isoform) under the KRT14 promoter in transgenic
mice produced a phenotype with supernumerary rudimentary mammary glands
(Mustonen T et. al, 2003; Mustonen T et. al, 2004; Figure 1.13F).

1.8.3.3 Msx1/Msx2 signaling
Msx1/Msx2 signaling is essential for the early mammary gland formation.
Developing mammary glands in Msx1/Msx2 double null mice fail to reach the
bud stage and further regress (Satokata I et. al., 2000; Satoh K et. al., 2004).

1.8.3.4 Parathyroid hormone-like peptide signaling
Normal signaling of the parathyroid hormone-like peptide (Pthlh, synonym:
Pthrp) through its cognitive parathyroid hormone receptor 1 (Pthr1) is required
for mammary gland progression through the primary sprout stage (Kobayashi T
et. al, 2005). Pthrp is expressed in the epithelial mammary bud and Pthr1 in the
underlying mesenchyme (Dunbar, ME et. al., 1999). Similar to humans with a
rare spontaneous loss-of-function mutation in PTHR1, mice with a targeted
deletion of Pthrp or Pthr1 lack mammary glands and nipples (Wysolmerski, JJ et.
al, 1998; Figure 1.13C). It was established that Pthrp controls the formation of
primary mammary mesenchyme that, in turn, maintains mammary fate of the
epithelial bud and, later, induces the nipple sheath. Unlike complete abolishment
of Pthrp signaling, its partial downregulation through regulated Pthr1 expression

107
allowed mammary gland development, but caused formation of abnormally small
nipples with altered morphology and significantly reduced smooth muscle.
Nipple abnormalities prevented proper milk flow and nursing of pups by mothers
(Kobayashi T et. al, 2005). Overexpression of Pthrp under the KRT14 promoter
results in expansion of the mammary mesenchyme in the ventral skin and
secondary induced nipple-like differentiation of the adjacent epidermis. The
above mentioned Pthrp null and Pthr1 null mice also have a poorly differentiated
fat pad precursor (Abdalkhani A et. al., 2002).

1.8.3.5 Tbx3 signaling
As a transcription factor Tbx3 is involved in the very early development of
mammary glands and it appears to act upstream of the Wnt pathway (Chapman
DL et. al., 1996). Mutant mice deficient for functional Tbx3 (Tbx3
tm1Pa
/Tbx3
tm1Pa

mice) do not form mammary glands (Figure 1.13E). No mammary glands form in
humans with ulnar mammary syndrome whose TBX3 gene carries a spontaneous
loss-of-function mutation (Rowley M et. al, 2004; Davenport TG et. al., 2003;
Papaioannou VE and Silver LM, 1998). Emerging data suggest that Tbx3
expression during early mammary gland development is positively regulated by
both Wnt and Fgf signalling through Fgfr1 (Eblaghie MC et. al., 2004).

1.8.3.6 Wnt signaling
Wnt signaling is believed to play key role in early induction. In TOPGAL
transgenic mice (STOCK Tg(Fos-lacZ)34Efu mice) carrying a beta-galactosidase
gene under the Wnt-responsive element, reporter activity is seen in the

108
presumptive milk line epithelium as early as E10.5 and is further confined to the
mammary placodes (DasGupta R and Fuchs E, 1999; Figure 1.13D). Early
expression of Wnt10b (previously: Wnt12), Lef1 and other members of the Wnt
canonical pathway compliment TOPGAL expression data (Christiansen, JH et.
al., 1995; Van Genderen C et. al., 1994). Abrogation of the Wnt signaling in the
Lef1 null mice causes developmental arrest at the early bud stage (Van Genderen
C et. al., 1994). Accordantly, overexpression of the Dkk1 (Dickkopf-1, WNT
inhibitor) in the skin under the KRT14 promoter prevents mammary gland
progression past the bud stage (Andl T et. al., 2002). Overexpression of Dkk1 in
the epidermis under the control of tetracycline/doxycycline-responsive promoter
in K5-rtTA, tetO-Dkk1, TOPGAL mice completely block mammary placodes
formation (Chu EY et. al, 2004; Figure 1.13D).

1.8.4 Sexual dimorphism of mammary glands in mice
As the part of the mouse sexual dimorphism in male embryos, mammary buds
and mammary mesenchyme degenerate between E13.5-E15.5 in response to
androgen signaling. Androgen receptors expressed in the mammary mesenchyme
is viewed as the direct target of testosterone (Kratochwil K, 1977; Dunbar M.E,
1999). Mammogenesis in the male mouse along with testicular feminization
(Ar
Tfm
) embryos deficient for the androgen receptor, proceed as it normally does
in females (Kratochwil K and Schwartz P, 1976).

109
Chapter 1: Morphoregulation of Ectodermal Organs.
Integument Pathology and Phenotypic Variations in K14-Noggin Engineered
Mice through Modulation of BMP Pathway.

Summary
Ectodermal organs are composed of keratinocytes organized in different ways
during induction, morphogenesis, differentiation, and regenerative stages. We
hypothesize that an imbalance of fundamental signaling pathways should affect
multiple ectodermal organs in a spatio-temporal-dependent manner. We produced
a K14-Noggin transgenic mouse to modulate bone morphogenic protein (BMP)
activity and test the extent of this hypothesis. We observed thickened skin
epidermis, increased hair density, altered hair types, faster anagen re-entry, and
formation of compound vibrissa follicles. The eyelid opening was smaller and
ectopic cilia formed at the expense of Meibomian glands. In the distal limb, there
were agenesis and hyperpigmentation of claws, interdigital webbing, reduced
footpads, and trans-differentiation of sweat glands into hairs. The size of external
genitalia increased in both sexes, but they remained fertile. We conclude that
modulation of BMP activity can affect the number of ectodermal organs by
acting during induction stages, influence the size and shape by acting during
morphogenesis stages, change phenotypes by acting during differentiation stages,
and facilitate new growth by acting during regeneration stages. Therefore during
organogenesis, BMP antagonists can produce a spectrum of phenotypes in a
stage-dependent manner by adjusting the level of BMP activity. The distinction
between phenotypic variations and pathological changes is discussed.

110
2.1 Introduction
The integument forms the interface between an organism and its

environment.
During development and evolution, different types

of epithelial organs form on
the body surface to allow animals

to adapt to different environments. Although
these organs, such

as hairs, glands, teeth, and so forth, appear to be very different

in structure and function, developmental studies suggested that

they are all
products of epithelial-mesenchymal interactions,

with variations overlaid on a
common theme (Chuong C-M, 1998). Genes involved

in human ectodermal
dysplasias have recently been cloned. These

studies show that a single gene
defect can cause abnormalities

in several ectodermal organs (Wisniewski et al.,
2002; Headon et al., 1999; Brunner et al., 2002). This further substantiates

the
notion that fundamental molecular pathways are frequently

shared in the building
of different epithelial organs. The commonly

used molecular pathways include
bone morphogenic protein (BMP),

fibroblast growth factor (FGF), sonic
hedgehog (SHH), Wnt, Notch,

Eda pathways and so forth (Chuong, et al., 2000a;
Fuchs et al., 2001). In each pathway there are multiple

ligands, receptors,
intracellular signaling transducers, and

extracellular antagonists. Knowledge of
these pathways motivates

us to investigate how these molecular activities are
translated

into tissue morphogenesis. In the context of tissue engineering,

such
knowledge will also be required to guide epithelial stem

cells appropriately to
form the tissues/organs desired.

Here we selected the BMP pathway for further analysis (Botchkarev et al., 2003).
BMPs

play an important role in many developmental systems. Initially

identified

111
for their effects on osteocyte proliferation and

differentiation, BMPs were further
shown to act as regulators

of proliferation, differentiation, apoptosis, cell
adhesion,

and migration during the development of multiple organs in many

organisms studied (Reddi et al., 1998; Miyazono et al., 2001; Wozney et al.,
2002). Loss-of-function mutations of various

components of the BMP pathway
lead to severe developmental abnormalities

often resulting in early embryonic
lethality (Itoh et al., 2000). The effect

of BMPs on proliferation, differentiation,
and apoptosis in

different developmental systems is complex. It is, however,

concentration-dependent. Low or high dosages of BMPs often result

in opposite
cell fate decisions: either proliferation or apoptosis (Piscione et al., 2001). BMP
activity in a given tissue depends on the concentration

and distribution of BMPs
and their antagonists. A number of

secreted proteins including Noggin, Follistatin,
Chordin, and

others antagonize BMP-mediated signaling (Massague et al., 2000).

Noggin is the most

powerful BMP-2 and BMP-4 antagonist (Zimmerman et al.,
1996; Piccolo et al., 1997). Different effects

of BMPs are often mediated by
distinct BMP receptors (BMPRs).

Several models suggest that the proliferative
effect of BMPs

is mainly mediated via BMP receptor IA (BMPR-IA; Yamaguchi
et al., 1999; David et al., 2001). Apoptotic

signaling is mainly mediated through
the receptor BMPR-IB (Kimura et al., 2000; Zhang et al., 1996). BMP signaling
is used in skin and skin appendages development.

In the presence of BMP-4,
ectodermal cells choose an epidermal

over a neural fate early in gastrulation
(Wilson et al., 1995). Later in skin development,

distinct spatial distributions of
different BMPs and BMPRs are

seen. BMP-6 and BMP-7 are mainly expressed in
the epidermis,

with BMP-7 present in the basal and BMP-6 in suprabasal layers

112
(Lyons et al., 1989; Wall et al., 1993; Takahashi et al., 1996). Also, BMPR-IA is
expressed in the basal layer, whereas BMPR-IB

is in the suprabasal layer. Based
on several lines of evidence

it was proposed that BMPR-IA mediates proliferation
effects

and BMPR-IB mediates differentiation effects of BMPs in epidermis
(David et al., 2001). Unlike BMP-6 and BMP-7, BMP-2 and BMP-4 are
expressed during

hair follicle (HF) organogenesis. BMP-4 is expressed transiently

in the mesenchymal condensations just before HF formation. Therefore,

this may
be part of the initial dermal signals inducing follicular

germ formation. BMP-2,
however, is expressed in the epidermal

placode, and in more advanced follicles it
is found in the matrix

and precortex cells (Bitgood et al., 1995; Botchkarev et al.,
1999). At the time of HF induction, BMP

signaling inhibits induction whereas
Noggin signaling stimulates

induction of HFs (Botchkarev et al., 1999;
Noramly,S. et al., 1998). Importantly, induction of secondary

(nontylotrich) HFs,
but not primary (tylotrich) HFs is affected

by BMPs/Noggin (Botchkarev et al.,
2002).

The role of Noggin during HF induction was addressed in the

Noggin
knockout mouse model. Data were provided usingNoggin

knockout skin grafts
because homozygous Noggin knockout mice

die prematurely. It suggests that
Noggin is important for secondary

HF induction. In this model, secondary HFs
failed to form. Induction

of primary HFs was not affected, yet their growth was
further

arrested because of long-term BMP excess. Similar to this, transgenic

mice engineered to overexpress BMP-4 in the outer root sheath

under the control
of the bovine cytokeratin IV promoter had

a complete deficiency of hair growth
after the first hair cycle

and, therefore, were progressively balding (Jamora et al.,
2003). It seems that

during development, Noggin prevents interactions between

113
BMPR-IA

and BMPs produced by the mesenchyme and placode. In support

of
this idea, Noggin treatment increases the hair placodes and

accelerates HF
morphogenesis in embryonic skin organ culture. Noggin is also required for HF
growth during postnatal life.

Normally, in adult HFs, Noggin activity is localized
to the

HF bulb. Noggin, produced by the dermal papilla, supports proliferation

in
the lower hair matrix. Overexpression of Noggin in proliferating

hair matrix cells
and differentiating hair precursor cells under

the proximal Mxs2 promoter leads
to the disruption of differentiation

in epithelial cells controlled here, in part, by
BMPs (Zhou et al., 1995). Another

important pathway in hair morphogenesis is
Wnt/ß-catenin

signaling and its up-regulation leads to an induction of excessive

numbers of HFs (Gat et al., 1998; Huelsken et al., 2001). Disruption of the
Wnt/ß-catenin

pathway in skin leads to an arrest of HF development (Andl et al.,
2002; Blessing et al., 1993). Inhibition of BMP activity is shown to produce
Lef-1 required

for the activation of ß-catenin/Lef-1 transcriptional

complex
(Botchkarev et al., 2001). Although the roles of Noggin in HF formation have
been studied,

its role in other skin appendages remains mostly unknown.
To

address these questions we created a transgenic mouse model

in which ectopic
Noggin expression was directed by the K14 promoter.

The K14-Noggin mice
study showed that Noggin mediates disruption

of normal BMP signaling during
development, causing multiple

abnormalities in a variety of ectodermal organs.
Hyperplasia

of pelage HFs occurred, ectopic HFs formed on the ventral side

of
the paw, supernumerary cilia formed in eyelids, and compound

vibrissa follicles
arose. Claws and footpads failed to form

normally. There were also defects in

114
organs that we do not normally

consider as skin appendages. For example, we
found an increase

in the sizes of external genitalia and defects in eyelid opening.

Here, we will describe an array of abnormalities and discuss

the roles of BMP
activity in pathogenesis.

2.2 Materials and Methods
2.2.1 Production and genotyping of transgenic mice
Mice were generated in the Norris Cancer Center transgenic mouse facility at the
University of Southern California. To generate transgenic mice, the inserts of
human keratin K14 promoter (Vassar et al., 1991) + chicken Noggin were
purified and microinjected into the male pronucleus of fertilized egg of C57BL/6J
x CBA/J mice followed by reimplantation of injected eggs into pseudopregnant
C57BL/6J x CBA/J females. The purification and microinjection of DNA were
performed as described (Liu et al., 1994). Mice were screened for transgene
presence by polymerase chain reaction (PCR) using chicken Noggin construct
specific primers: (5-CCAGATCTATGGATCATTCCCAGTGC-3 and
5-GGAGATCTCTAGCAGGAGCACTTGCA-3). Tail genomic DNA was
extracted as described (QIAGEN). PCR products were amplified in separate
reactions using the three stage PCR program: 94
o
C for 2 min; 94
o
C for 1 min,
55
o
C for 1 min, 72
o
C for 1 min (30 cycles); 72
o
C for 10 min. The identities of
the founder mice were confirmed by Southern blot analyses.

115
2.2.2 Quantitative genotyping
Quantitative genotyping of K14-Noggin mice was done by real-time qPCR using
SYBR Green technology (Dhar, et al., 2001). The reaction and detection were
carried out in GeneAmp 5700 (PE Applied Biosystems, Foster City, CA, User
Manual). A separate set of primers was designed for chicken Noggin to be used
for real-time qPCR: (5-TCTGTCCCAGAAGGCATGGT-3 and
5-CGCCACCTCAGGATCGTTAA-3). To control differences in the quantity of
DNA template, the mouse L-32 gene was amplified in parallel for each sample
and was used as a normalization factor to calculate the relative amount of chicken
Noggin in mouse genomic DNA. Following L-32 specific primers were used:
5'-TGGTTTTCTTGTTGCTCCCATA-3'
and 5'-GGGTGCGGAGAAGGTTCAA-3'. A detailed protocol for real-time
qPCR is described elsewhere (Hizer et al., 2002). Among all K14-Noggin mice
tested we selected one mouse that showed the highest dC
T
value (5.5) on
real-time qPCR (C
T
- cycle threshold value, dC
T =
C
T
of Noggin - C
T
of L32 for
the same sample). This mouse contained the lowest number of K14-Noggin in the
genome. Relative amount of K14-Noggin in all other mice was calculated as
following: Fold difference = 2 x (5.5 - dC
T
).

2.2.3 Quantitative RT-PCR
Amount of chicken Noggin mRNA in K14-Noggin mice tissue was measured
using real-time quantitative RT-PCR method, based on SYBR Green technology,
mentioned above. RNA was extracted from the mice ear pinna using RNeasy
Mini Kit, following manufacture protocol (QIAGEN). Ear pinna were selected

116
for this experiment because they contain two layers of K14 expressing epidermis
within relatively small amount of tissue. Real-time PCR was carried out using
identical set of primers and following the same protocol as for the quantitative
genotyping described in the previous paragraph. Relative amount of Noggin
mRNA in mice tissue was calculated as following: Fold difference = 2 x (9.45 -
dC
T
), where 9.45 is the dC
T
value for the mouse with the lowest level of chicken
Noggin mRNA in the tissue.

2.2.4 Histological, histochemical and immunohistological staining
Tissues were collected and fixed in 4% paraformaldehyde in phosphate-buffered
saline (PBS), dehydrated, embedded in paraffin and sectioned at 5-6 um. When
necessary, specimens were additionally decalcified in Immunocal solution for 48
hours at 4
o
C (American Master*tech Scientific) after fixation. Standard H&E
staining was performed for basic histological analysis. Frozen tissue sections
were used for the tyramide based tyrosinase assay (TTA; Han et al., 2002).
Briefly, after 3% peroxide treatment and 5% BSA (fraction V) blocking, the
avidin/biotin blocking was performed (Vector lab). Next, tyramide-biotin in 1x
application diluent was applied (PerkinElmer Life Sciences). Following a
washing step, Streptavidin-CY3 was applied (1:600, Sigma). Immunostaining
was performed using the Ventana Discovery automated immunostaining module
(Ventana Medical Systems
TM
). Primary antibodies used were mouse monoclonal
anti-PCNA (1:500, Chemicon), rabbit anti-K14 (1:400, Berkeley Antibody
Company);

anti-K10 (1:200, Sigma). DAB detection kit (Ventana Medical
Systems
TM
) was used for color development.

117
2.2.5 In situ hybridization
Mouse tissues from various ages were used for section in situ hybridization.
Section in situ samples were fixed and dehydrated according to the standard
protocol. All solutions used for the procedure were DEPC-treated to inactivate
RNAse. To detect the RNA expression, the tissue was hybridized with
digoxigenin-labeled probes. The signals were detected by using an
anti-digoxigenin antibody coupled to alkaline phosphatase. Some tissue samples
were processed using the Discovery automated in situ hybridization instrument
(Ventana Medical Systems
TM
). Whole mount in situ procedure was performed on
E15 mouse embryos. Specimens were fixed in 4% paraformaldehyde in
DEPC-treated PBS. Tissue samples were then dehydrated in a series of methanol
in PBS + 0.1% Tween 20 (PBT buffer) and stored in absolute methanol at 20°C
before the actual staining procedure. Whole mount in situ hybridization
procedures were performed using the InsituPro automated In Situ detection
module (Intavis AG). Analysis was performed according to the standard whole
mount in situ protocol (Jiang et al., 1999).

2.2.6 Morphometric analysis
All surgical procedures were performed on anaesthetized mice (Ketamine
HCl:Xylazyne mixture was used). For the whole mount skin preparation anagen
skin was collected, inverted and subcutaneous tissue was removed. These
samples were fixed and dehydrated in a stretched condition. After dehydration,
skin samples were cleared with xylene and photographed. Morphometric analyses
were then performed using Adobe PhotoShop. Analysis of hair shaft structure

118
was performed under the inverted microscope according to previously described
protocol (Millar et al., 1999; Nakamura et al., 2001). Relative number of guard,
awl, auchene and zigzag hairs was determined in the fur of the dorsal skin.
Length of the hair growth cycles stages was measured in mice while they were
two to six months old. Length of the anagen and telogen stages of the hair growth
cycle was established based on the change of the skin color from pink during
telogen to black during anagen. These changes occur due to the active
melanogenesis in the hair follicles during anagen and were proven to be valid
criteria for the hair cycle staging elsewhere (Slominski, et al., 1994). All
observations were performed on shaved mice. External genital measurements
were performed on anesthetized animals. Non-erect genitalia were measured in
both control and K14-Noggin animals.

2.2.7 Scanning electron microscopy (SEM) analysis
Tissues were prepared according to the standard SEM protocol. Briefly, it
includes fixation in 2.5% glutaraldehyde in 0.1 M sodium cacodylate,
dehydration, and critical point drying from ethanol. Next, samples were coated
with gold in a sputter coat chamber. They were examined by SEM.

2.3 Results
2.3.1 Production, genotyping and phenotyping of K14-Noggin mice
The chicken Noggin cDNA fragment was subcloned into the human

K14 vector
(Figure 2.1A). K14-Noggin-PolyA inserts were released

from plasmids and used
for injection. Three independent transgenic

lines were produced with similar

119
Figure 2.1 Production of K14-Noggin mouse.
A: K14 Noggin construct used to generate transgenic mouse. The size of insert
used and restriction enzyme are indicated. B: Genotyping of K14-Noggin mutant
mouse. Products of PCR reaction using specific primers (see Materials and
Methods for primers used). Lane 1, DNA standard; lane 2, positive
controlK14-Noggin founder mouse; lanes 3 and 6, wild-type mice; lanes 4 and
5, K14-Noggin-positive mice. C: Appearance of control C57BL/6J (left) and
mutant K14-Noggin 2.5-month-old mice. Note the obvious hypertrichosis of the
K14-Noggin mouse. The eye opening is small (white arrow). Digits are not
distinctly separated from each other. Regions to be studied further in each figure
are marked by red brackets.

120
Figure 2.1: Continued

121
phenotypes. Identities of K14-Noggin

mice were confirmed by PCR-genotyping
using chicken Noggin-specific

primers (Figure 2.1B). We have isolated several
lines and found

a range of phenotypes. There were mice with severe phenotypic

changes (Figure 2.1C) and mice with moderate alterations. The

more severe
phenotypes included absence of claws, interdigital

webbing of the paws,
hypertrichosis all over the body, shortening

of the telogen period of the hair
growth cycle, and increased

size of the genitalia. In other mice pathological
changes were

milder. We performed quantitative genotyping of the K14-Noggin

mice and quantitative RT-PCR for the Noggin mRNA to establish

whether
strength of phenotypic changes correlates with K14-Noggin

transgene copy
number in the mouse genome and Noggin expression

level. We studied several
key phenotypic features of the limbs

of every mouse and correlated them with the
fold difference

values (see Materials and Methods). Based on this we divided

all
K14-Noggin mice into low-transgenic (TG) copy number and

high-TG copy
number with fold difference value for high-TG copy

number animals equal to 3
and higher.

2.3.2 Phenotypes in the head
Eyelids. Adult high-TG copy number K14-Noggin mice had smaller eye
openings

(Figure 2.2A and 2.2B), whereas the diameters of the eyeballs

were not
significantly different (3.9 ± 0.5 mm in control

mice and 3.75 ± 0.05 mm in
high-TG copy number K14-Noggin

mice, n = 3). Smaller eyelid openings were
already obvious as

early as postnatal day 14.

122
Figure 2.2 Pathology in eyelid, an epithelial appendage to protect the eye. A:
Normal size eye slits of 4-month-old control C57BL/6J mouse. B: Narrow eye
slits in 4-month-old K14-Noggin mouse. Eyelids remain partially fused. Eyeballs
are of the same size (not shown). Arrowheads point at inward-growing cilia. CF:
Eyelid suture of newborn control mouse. C: Strong K14 immunostaining is seen
in the epidermal side, suture, and conjunctival side of the eyelid. DF: Whole
mount in situ hybridization of 1-mm-thick eyelid stripe. Green color is computer
pseudo-coloring. BMP2 transcripts are expressed predominantly on the
conjunctivae side whereas BMP4 is distributed more evenly throughout the
eyelid suture. Msx2 transcript distribution is similar to BMP4. G and H: Corner
of eyelids (commissure, corresponding to the areas outlined with red on A and B)
of adult mice. Formation of ectopic pillosebaceous units at the expense of
Meibomian glands in high copy number TG K14-Noggin mice (hcn TG
K14-Noggin mice). Scale bars, 100 µm.

123
However, no significant delay of

eyelid opening was seen in K14-Noggin mice in
comparison with

the control mice. In addition, adult eyelids of high-TG copy

number K14-Noggin mice exhibited abnormalities. Most significantly,

there was
formation of ectopic pilosebaceous units at the expense

of Meibomian glands.
Extra cilia grew in different directions,

often pointing inwards toward the cornea,
resulting in entropion

[Figure 2.2; B (inset) and G and H (arrows point at
inward-growing

cilia)]. No prominent corneal lesions were noticed. We examined
the eyelids of newborn mice. Immunohistochemically

K14 protein expression
was detected in the outer epidermis,

conjunctival epithelium, and eye suture of
both upper and lower

eyelids (Figure 2.2C). The observed suture defects are
consistent

with the observation that BMPs are expressed in the eye suture

of
control E15 mouse embryos. Both BMP4 and BMP2 are expressed

in the eye
suture epithelium with BMP2 specifically at the conjunctival

side and BMP4
throughout the length of the epithelial suture

(Figure 2.2, D and E; these are thick
preparations with whole

mount in situ hybridization). Msx2 (downstream of
BMPs in other

systems) is also expressed in the eye suture (Figure 2.2F).
Vibrissae. Vibrissae are specialized, highly innervated HFs with somatosensory

function. Normal mice have one vibrissa emerging from one individual

vibrissae
follicle, which is enclosed in a collagen capsule

(Figure 2.3, A and C). In
high-TG copy number K14-Noggin mice,

examination showed that several
vibrissae emerged from one orifice

(Figure 2.3B). Histological sections showed
that two or three

combined vibrissae follicles shared one capsule (Figure 2.3, D

and E). Histological sections showed these compound follicles

share a common
upper part

124
Figure 2.3 Pathology in vibrissae and pelage hairs.
A and B: Control and K14-Noggin vibrissae HFs. K14-Noggin mice have
compound follicles that share one orifice and one capsule. The number in B
indicates the number of vibrissa filaments that share the same orifice. For
example, 1 + 2 means one normal follicle with one filament from one orifice plus
one compound follicle with two filaments growing from one orifice. CE: H&E
stain of control (C) and K14-Noggin (D, E) vibrissae HFs. K14-Noggin follicles
share part of the same outer root sheath, open into the same canal but have a
distinct dermal papillae and matrix, and produce a separate inner root sheath and
fiber. F and G: View of inverted skin from the dorsal trunk region of the control
and K14-Noggin mice. In the control mouse, all HFs are in anagen. Distinct
primary (big) and secondary (small) HFs can be identified. In the K14-Noggin
mouse, the density of HFs is increased and the difference between primary and
secondary follicles is not obvious. H: Density of HFs per 3 mm2 in control (blue)
and K14-Noggin (red) mice. Density is increased by ~80% in the K14-Noggin
mouse in comparison with the control mouse. I: Relative size distribution of HFs
in control (blue) and K14-Noggin (red) mice (size corresponds to the diameter of
the hair bulb; see inset in Figure 4F ). In the control mouse, HFs clearly
fractionate into two distinct groups: those with smaller size (primarily secondary
HFs) and those with larger size (primary HFs). In the K14-Noggin mouse, there
is no clear fractionation of HFs. The majority of HFs are of intermediate size.
This could be because of increased proportion of secondary awl and auchene
hairs (see Results). J and K: H&E staining of skin sections from the back of the
control and

125
Figure 2.3: Continued
K14-Noggin 2-week-old mice. Normal secondary and primary (bottom) HFs are
seen in the control mouse. In some regions of the K14-Noggin mouse, there are
enlarged HFs and hypertrophic sebaceous glands. Some hair fibers point to
wrong directions. Spacing between follicles is reduced. Skin epidermis is
thickened. L and M: H&E staining of longitudinal sections of the tail from the
control and K14-Noggin mice. Different sizes of hypertrophic HFs pointing in
different directions are seen in the K14-Noggin mouse. The total number of
follicles has increased. Some follicles are dystrophic and appear to have multiple
dermal papillae. The epidermal and dermal layers appear thicker. Scale bars: 1
mm (B); 100 µm (CE and JM).

126
Figure 2.3: Continued

127
Figure 2.3: Continued

128
of the outer root sheath, but have

separate lower follicle regions with independent
dermal papilla

and matrix. They produce separate inner root sheath and vibrissa

fibers that open into the same canal. Because they seem to derive

from the shared
outer root sheath, we consider them to belong

to the category of compound HFs
(Oro et al., 2003).

2.3.3 Phenotypes on the trunk
Pelage hair. On the whole body, K14-Noggin mice exhibited a prominent
hypertrichosis.

The hypertrichosis observed here is a combined result of increased

hair density and an excessive mass of hair filaments. Whole

mount morphometric
analysis of anagen skin from control and

K14-Noggin mice was done to quantify
hair density and size distribution

of HFs (Figure 2.3, F and G). The wild-type
mice hair density

was 35.1 ± 2.5 HFs per mm
2
. In the high-TG copy number

K14-Noggin mice hair density was 63.4 ± 3.9 HFs per mm
2
,

80% higher than
the control mouse (Figure 2.3H). The low-TG copy

number K14-Noggin mice
hair density was 50.0 ± 7.3 HFs

per mm
2
, 42% higher than the control mouse.
The maximal width of the follicles was then measured as an indication

of follicle
size (Figure 2.3F, inset, red arrows). In the control

mice, all HFs are clearly
distributed into two distinct groups:

smaller size HFs (secondary HFs) and larger
size HFs (primary,

guard HFs). In contrast, in high-TG copy number K14-Noggin
mice

there was no clear fractionationing of the HF sizes (Figure

2.3I). The
majority of HFs were of smaller size, probably representing

secondary follicles.
Analysis of hair shaft structure showed

that high-TG copy number K14-Noggin
mice fur contains all four

types of hair fibers: guard, awl, auchene, and zigzag.

129
However,

high-TG copy number K14-Noggin mice have more awl and auchene

hairs than wild-type mice. Our control mice showed 64.3 ±

6% of zigzag hairs
and only 27.9 ± 3.6% of awl and auchene

hairs, high-TG copy number
K14-Noggin mice have 51 ±

4.5% of zigzag and 43.4 ± 3.8% of awl and auchene
hairs.

We believe that the excessive number of awl and auchene hairs

is mostly
responsible for the increase in hair density in K14-Noggin

mice. Awl and
auchene hairs are generally larger than zigzag

hairs and this may result in the
larger proportion of intermediate

sizes of HFs (Figure 2.3I). Guard hairs are
present and appear

to have no significant change in their number in K14-Noggin

mice. On the sections of the dorsal trunk skin, control skin contained

primary and
secondary HFs (Figure 2.3J). In the K14-Noggin mouse,

there were regions of
skin with secondary HFs appearing to be

normal (not shown) and regions of skin
with abnormally enlarged

HFs (Figure 2.3K). There are hypertrophic sebaceous
glands and

randomly oriented hair fibers. The epidermis of interfollicular

skin is
thickened. On the sections of the tail, the control

skin exhibited secondary HFs
and tail scales arranged in a regular

pattern (Figure 2.3L). In the K14-Noggin tail,
some regions were

relatively normal (not shown) but some regions showed
drastic

pathology: cystrophic HFs of different sizes pointing to different

directions (Figure 2.3M). Some follicles appeared to have multiple

dermal
papillae. The density of HFs increased. The dermal layer

appears to thicken at the
expense of the adipose layer. Whether

this is a direct or indirect systematic effect
remains to be

determined. We measured the length of the anagen and telogen of
the hair

growth cycle in high-TG copy number K14-Noggin mice. On average,

their anagen length was 12.3 ± 1 day, and telogen length

was 7.6 ± 2 days. In

130
control mice the anagen length was

not significantly different (13.4 ± 1 day), but
the telogen

length was significantly greater, ranging from 12 to 40 days.
Compared to wild-type mice, K14-Noggin mice showed distinct

differences in
their external genitalia. Overall, the sizes

of both male and female external
genitalia were bigger than

those in the control animals (Figure 2.4; A to D). When
freed

from the prepuce, the distal segment of the os penis in the

K14-Noggin male
mice was significantly longer than that of the

control mice (Figure 2.4, F and G).
During embryonic development

at E15, the tip of growing glans penis expressed
high levels

of BMP4 (Figure 2.4E). Histologically, the preputial lamella was
significantly thickened

in the K14-Noggin mice. At the same time, the
differentiation

of hairy spines, mechano-sensory structures of the preputial

lamellae, was suppressed. Hairy spines are periodically arranged

skin appendages
composed of both epidermal and dermal components.

They start to differentiate at
postnatal week 1 and undergo

keratinization at week 2 (Figure 2.4H). However,
in 2-week-old

high-TG copy number K14-Noggin mice, hairy spines remained
primarily

undifferentiated (Figure 2.4I). In situ hybridization showed

high levels
of BMP4 expression in the epithelial compartment

of all hairy spines (Figure
2.4J). Immunostaining showed strong

K14 keratin expression in the basal layers
of both the penis

and prepuce in the control and K14-Noggin mice (Figure 2.4, K

and L). K10 keratin expression showed that differentiation

of hairy spines in
2-week-old K14-Noggin mice was suppressed

(Figure 2.4, M and N). These mice
were fertile.

131
Figure 2.4 Pathology in external genitalia.
A and B: External genitalia of the control and K14-Noggin female mice. C and
D: External genitalia of the control and K14-Noggin male mice. E: BMP4
expression pattern in the external genitalia of E15 mouse embryo. Very strong
expression in the glans penis. F and G: Side-to-side comparison of the control
and K14-Noggin male external genital. H and I: Hairy spines of the 2-week-old
control and K14-Noggin mice. Hairy spines in the K14-Noggin mouse are not
completely developed. J: BMP4 expression in the developing hairy spine of the
2-week-old control mouse. KN: K14 and K10 keratin expression patterns in the
preputial lamella and developing hairy spines on the border of the glans and
prepuce from a 2-week-old wild-type and K14-Noggin mouse. Note weak K10
expression in the immature hairy spines of the K14-Noggin mouse (N). Scale
bars 25 µm.

132
Figure 2.4: Continued

133
Figure 2.4: Continued

134
2.3.4 Phenotypes in the distal limb region
Claw. During embryonic development at E15, BMP4 is specifically expressed

in
the mesenchyme at the tips of the digitsthe sites

of claw formation (Figure
2.5A). K14 keratin is highly expressed

in the claw matrix (Figure 2.5B) and
overexpression of Noggin

in these regions could potentially perturb claw
development.

Indeed, a prominent feature of K14-Noggin mice was the claw

phenotype. It ranged from split claws in low-TG copy number

mice to complete
absence in high-TG copy number mice (Figure

2.5; E to H). Absence of claws in
high-TG copy number K14-Noggin

mice was coupled with polydactyly and
interdigital webbing (Figure

2.6B; Moore et al., 1998). In the wild-type claw,
highly proliferative epithelial cells

are confined to one region, known as the claw
matrix (Figure

2.5C). In the low-TG copy number K14-Noggin mice, the claw
matrix

was not distinct and PCNA-positive cells were not confined to

one place,
but rather distributed widely in the whole claw region

(Figure 2.5D). This can
account for the multiple growth centers

and multiple keratinized plates within one
claw. Claw plates

apparently grew parallel and were separated from each other.

Patches of epidermis often separated one plate from another

(Figure 2.5G). In
high-TG copy number mutant mice, all claws

on forelimbs and almost all claws
on hindlimbs were replaced

with a thickened cornifying epidermis (Figure 2.5, H
and J). In control mice, the epidermal differentiation marker K10 keratin

is found
only in the hyponichium and nail fold, but not in the

claw itself (not shown). In

135
Figure 2.5 Pathology in claws.
A: E15 wild-type mouse paw in situ with BMP4 probe. White arrows point to
expression sites at the tips of the digits. Black arrow points to the group of
expression sites corresponding to footpads. BMP4 expression in the mesenchyme
of the developing claw is shown on the inset. B: Strong K14 expression in
epithelial cells of proliferating claw matrix (green box) and beyond it. C and D:
Expansion of proliferating area in the claw of low-TG copy number K14-Noggin
mice (lcn TG K14-Noggin mice). PCNA-positive cells are limited to the claw
matrix (green box) in wild-type mice, but extend all of the way to the tip of the
digit in K14-Noggin mice (green boxes). There are multiple proliferating centers
within one claw of the K14-Noggin mice. E to H: Progressive abnormalities in
the claw of the K14-Noggin mice. Compared to the control claw from C57BL/6J
mouse (E), claws in the low-TG copy number K14-Noggin mice split into
separate plates (F) and are partially substituted by epidermis (G, demarcated with
a green dotted line). No claw is identifiable in the high-TG copy number
K14-Noggin mice (H, white arrows). I and J: K10 keratin expression in low (I)
and high (J) TG copy number K14-Noggin mice. In low-TG copy number mice
claws are K10-negative (arrows) despite the apparent macroscopic and
microscopic abnormalities (see above). In the high-TG copy number mouse the
epidermis that substitutes the claw expresses K10 (arrows). In the control mouse
(K, inset) the claw is K10-negative. K and L: Pigmentation of the claw in low-TG
copy number K14-Noggin mice. Abundant pigment deposits in the claw
epithelium (L). Tyrosinase activities (red color, K) associated with pigment
production are seen.

136
Figure 2.5: Continued

137
Figure 2.5: Continued

138
Figure 2.5: Continued

139
Figure 2.6 Pathology of the ventral side of the paw.
A and B: Normal and hypoplastic foot pads from the wild-type and K14-Noggin
mouse, respectively (white and black arrows). Presence of additional digit and
interdigital webbing are marked in K14-Noggin mouse paw. C and D: Scanning
EM normal glabrous skin and skin with ectopic HFs on the ventral side of the
digits from wild-type and K14-Noggin mice, respectively. E: BMP4 expression
in the mesenchyme of the developing footpad in E15 control mouse embryo. F
and G: Suprabasal expression of BMP2 in the epidermis of the adult footpad (F)
and on the border of the footpad (G). Note the sharp decline of BMP2 expression
on the border of the footpad. H to J: Growing footpad of the newborn mouse.
Active proliferation is seen in the mesenchymal condensation of the footpad as
judged by PCNA staining (J). Eccrine glands are forming at this time (marked by
arrowheads at H). K14 is expressed in the basal layer of the footpad epidermis (I).
K and L: H&E of the footpad from control and K14-Noggin mice. Typical
afollicular epidermis and eccrine glands are seen in the control mouse (K). In the
K14-Noggin mouse, multiple HFs replace the eccrine glands. M and N: H&E of
the skin on the digit from K14-Noggin mouse. M: Tip of the digit. Claw is absent.
It is substituted by a hyperplastic patch of epidermis with sebaceous glands and
HFs. N: Ventral side of the distal digit. Many HFs with sebaceous glands have
formed, instead of sweat glands and just a few HFs in the normal.

140
Figure 2.6: Continued

141
Figure 2.6: Continued

142
the high-TG copy number mutants,

the epidermal thickening expressed K10
keratin (Figure 2.5I).

Claws of the low-TG copy number K14-Noggin mice
differentiated

properly and were K10-negative. Patches of epidermis that
separated

multiple claw plates from each other were otherwise K10-positive

(Figure 2.5J).

Another interesting phenotype was the pigmentation of the claw.

Control mice
have black fur, but lack pigmented claws (Figure

2.5E, inset). In our K14-Noggin
mice, we have not seen changes

in fur color. However many claws of low-TG
copy number K14-Noggin

mice and all hindlimb claws of high-TG copy number
mutants were

hyperpigmented (Figure 2.5K, inset). On histological sections,

unlike in the control mice, basal and suprabasal layers of claws

from K14-Noggin
mice contained abundant melanin granules (Figure

2.5L). Tyrosinase activity, a
general marker of melanocytes,

was detected in the pigmented claws (Figure
2.5K).
Ventral Paw. There are several changes in the ventral paw integuments in

high-TG copy number and low-TG copy number K14-Noggin mice.

Normally,
there are six footpads (Figure 2.6A) that have evolved

for land habitats (Guha et
al., 2002). BMPs are specifically expressed in the

footpads. At E15, developing
footpad areas showed localized

BMP4 expression in the mesenchyme (Figure
2.6E). At later developmental

stages and in adults BMPs expression shifts to the
epidermis.

Both BMP2 and BMP4 are specifically expressed in the footpads

epidermis and are down-regulated in the epidermis outside of

the footpads
(Figure 2.6, F and G). K14 is expressed in the epidermis

of developing footpads

143
(Figure 2.6I). Localized proliferation

in the mesenchyme results in footpad
growth (Figure 2.6, H and

J). In high-TG copy number K14-Noggin mice,
footpads are hypoplastic

(Figure 2.6B). Footpads in mutant mice are markedly
shallower,

but the total number of footpads does not change.

Normally, the
ventral paw has glabrous skin and does not contain

any significant amount of
hairs (Figure 2.6C). Footpads are completely

devoid of hairs (Figure 2.6K). The
ventral paw skin contains

eccrine glands, which are particularly dense at the tip of
the

digits and in the footpads (Figure 2.6K). In contrast, numerous

HFs in
high-TG copy number K14-Noggin mice were found on the

ventral side of the
paws, including footpads where we observed

nearly complete substitution of
eccrine glands by HFs (Figure

2.6; L to N).

2.4 Discussion
The formation of ectodermal organs depends on a series of topological
transforming activities of the epithelial sheet such as folding, thickening,
branching, and so forth. These processes are based on local cellular behaviors
including proliferation, differentiation, re-arrangement, and apoptosis. The
formation of all ectodermal organs goes through induction, morphogenesis, and
differentiation stages. In addition, some ectodermal organs undergo
cycling/regeneration stages (Hamrick et al., 1998). They share fundamental
signaling pathways during these developmental stages and thus they are
variations overlaid on the common theme (Wu, et al., 2000; Chuong et al., 1999).
Serious defects of these basic signaling pathways tend to generate multiple

144
ectodermal organ defects as seen in some forms of ectodermal dysplasia
(Chuong, et al., 200b). Milder perturbation of the strength of these pathway
activities may lead to morphoregulation, i.e., the modulation of the
morphological phenotype of the organ (Slavkin et al., 1998). Here we further
propose that when the perturbation mainly acts during induction stages, the total
number of a given ectodermal organ is altered. In contrast, when the perturbation
mainly acts during morphogenesis stages, the size and shape of the organ may
change. Finally, when perturbation acts mainly during differentiation stages,
maturation of the organ is affected. We tried to modulate one of the molecular
pathways to evaluate the validity and the scope of this model. We analyzed the
multiple roles of the BMP pathway in ectodermal organ morphogenesis using
K14-Noggin mice. The onset of transcriptional activation of keratin 14 and its
partner keratin 5 was studied previously (Edelman et al., 1992). It was shown that
their transcription was first detected as early as E9.5. At early times, expression is
restricted to certain areas of the embryo, such as the ectoderm of the developing
facial structures. At E13.5 to E14.5, there is a dramatic increase and expansion of
K14 and K5 promoter activity in ectoderm throughout the body of the embryo.
This coincides or precedes development of the ectodermal organs affected in
K14-Noggin mice by overexpressed Noggin. However, the minimal
concentration of Noggin required to perturb a particular ectodermal organ can be
reached during induction, morphogenesis, or differentiation stages of that
particular organ, thus producing different phenotypes. In several occasions, rather
than producing serious

145
Figure 2.7 Summary of multiple epidermal organ defects caused by disruption of
BMP pathway in the skin. A: Defective regions are shown in gray shades. B:
Phenotypes are summarized. They are grouped based on the developmental
stages when defects occur.

146
pathological changes with functional impairment, Noggin altered ectodermal
organ number, size, and differentiation status. Here, we focus on the multiple
ectodermal organ defects caused by suppression of the BMP pathway (Figure
2.7), and discuss the concept of pathological changes and phenotypic variations.

2.4.1 BMP Regulates the Number, Size, Type, and Cycling of HFs
In the adult mouse, two types of hairs can be found: primary

(tylotrich) and
secondary (nontylotrich). Secondary hairs are

further classified into awl, auchene,
and zigzag based on the

shaft structure. Primary and secondary HFs start to
develop

at E14.5 and E16.5 accordingly during embryogenesis (Byrne et al., 1994;
Vielkind et al., 1996; Philpott et al., 1998), and

may depend on different
molecular pathways. Primary HF morphogenesis

is highly dependent on the Eda
pathway (Paus et al., 1999). Secondary HF morphogenesis

is highly dependent on
the level of BMPs and their antagonists,

such as Noggin. Using the
Noggin-knockout mouse model, it was

previously shown that excessive amounts
of BMP leads to the

inhibition of the secondary, but not primary HF formation.

Our results indicate that excess of Noggin affects hair induction

and results in an
increase in HF density by up to 80%. Primarily

the number of secondary awl and
auchene hairs are increased.

This is consistent with the idea that BMPs/Noggin
specifically

control and modulate secondary HF formation. Noggin may result

in
a higher density of HFs in two ways. First, by lowering the

threshold for the
induction of awl and auchene HFs, and second,

by lengthening the competence
period and extending the inductive

phase of these HFs further into postnatal life.

New HFs can be induced from interfollicular epidermis or from

the outer root

147
sheath of the pre-existing HFs, as in the case

of compound follicles. Compound
HF formation was observed

in K14- N87ßcat mice that express a stabilized form

of ß-catenin under the K14 promoter. In K14- N87ßcat

mice, supernumerary HF
formation takes place continuously throughout

postnatal life and is associated
with pilomatricoma formation

in the skin. In the K14-Noggin mice, induction of
new pelage

HFs primarily occurred in the interfollicular epidermis. However

in
vibrissae, we observed many compound follicles as they all

share one
infudibulum and outer root sheath. We propose that

the presence of Noggin
lowers the threshold for the induction

allowing additional HFs to be induced from
the interfollicular

epidermis or the outer root sheath. Using the same reasoning,

ectopic HFs were induced from the glaborous skin of the ventral

paws including
footpads. Previously, it was reported that adult

dermal papillae transplanted under
glabrous epidermis could

induce new HF formation (Laurikkala et al., 2002;
Jahoda et al., 1992). The adult dermal papilla is a

strong site of Noggin signaling
(Botchkarev et al., 1999), and Noggin may be responsible

in part for the inductive
abilities of the dermal papillae. Noggin shows a distinct spatio-temporal
distribution during

hair development. During development at E15.5 to E17.5
Noggin

is expressed in the mesenchyme underneath the epidermis. In

the adult
mouse, its expression is restricted to the dermal

papillae. Broad Noggin
expression at E15.5 to E17.5 in the

skin coincides with induction of nontylotrich
HFs. Mesenchymally

derived Noggin competes with BMP2, four ligands for
binding

to BMPR-IA in hair placodes. In our mouse model, excessive Noggin

produced in the epidermis under the K14 promoter strongly inhibited

BMP2 and
BMP4 signaling during the crucial time of HF formation.

Therefore, the

148
activator/inhibitor ratio in the microenvironment

was tilted toward the activator
resulting in additional HFs.

Hairs in K14-Noggin mice cycled significantly faster than in

control mice.
Although there was no significant change in the

length of the anagen, the telogen
in K14-Noggin mice was markedly

shorter than the telogen observed in control
mice. Our observation

is consistent with the effect of exogenous Noggin delivery
into

the telogen skin on wild-type mice. Implantation of beads soaked

with
Noggin results in hair growth induction (Jahoda et al., 1993). Our results

add to
the evidence that the BMP pathway is an important regulator

of the
telogen-anagen transition.

2.4.2 BMP Affects the Development of Claw and Integuments of the Distal
Limbs
In the paw of K14-Noggin mice, we observed syndactyly and postaxial

polydactyly, consistent with what was reported previously. In K14-Noggin mice,
the induction, morphogenesis, and differentiation

stages are affected. Claw
agenesis was seen in high-TG copy

number mice. The claw induction failed and
the claw fields underwent

alternative epidermal differentiation. Low-TG copy
number mice

developed aberrant claws. During the morphogenesis stage the

original claw field splits into several claw plates, but they

remain in the same
plane. At later differentiation stages, these

claws show abnormal differentiation

149
with loss of function. Claw

morphology and differentiation is affected because
claw matrix

cells are K14-positive. An excess of Noggin in the claw matrix

affects normal proliferation and delays claw-specific differentiation.

Localized
zone of proliferating cells normally located next

to the eponychium is expanded
and proliferating cells, in isolated

patches, are present all of the way toward the
tip of the claw

in K14-Noggin mice. It is interesting to note that claw
malformation

was also reported for the Msx2-Noggin transgenic mouse and

Msx2
is known to be expressed in the claw area (our data, not

shown). However the
Msx2-Noggin claw phenotype was not described

in full to allow comparison with
the K14-Noggin phenotype.

Hyperpigmentation of the claws is a prominent feature of K14-Noggin

mice. In
mouse skin, active melanogenesis occurs only in the

matrix of anagen HFs. Stem
cell factor is expressed in epithelial

cells of the hair matrix and is important for
stimulating melanogenesis (Botchkareva et al., 2001). If stem cell factor is
constitutively expressed under the K14

promoter, the epidermis becomes
pigmented (Nishimura et al., 2002). We speculate that

Noggin may up-regulate
the stem cell factor in the claw, thus

causing activation of melanogenesis.
Footpads displayed major modifications in the ventral paw. Localized
mesenchymal cell proliferation is a key event during

footpad development (Mori
et al., 2000). Epidermal thickening and eccrine sweat

gland formation
accompanies mesenchymal expansion. Excess of

Noggin results in hypoplastic
footpads. Noggin suppresses localized

mesenchymal cell proliferation that is
otherwise positively

controlled by BMPs. Noggin also suppresses eccrine sweat

150
glands

and causes formation of pilosebaceous units. Noggin may abrogate

the
induction of sweat glands and induce HFs as alternative

skin appendages, or they
may trans-differentiate the induced

early sweat gland primordia into hairs.

2.4.3 BMP Decreases the Size of Eyelid Opening
The morphogenetic process for the opening of the eyelids is

affected in
K14-Noggin mice. This leads to small eye openings

and abnormally shaped
eyelids, especially obvious in lateral

and medial commissures. Excessive Noggin
also suppresses induction

of Meibomian glands and induces formation of many
ectopic cilia

often pointing inwards toward the cornea. The extent of these

pathological changes is highly dependent on the transgene copy

number. Severe
eyelid abnormalities were seen only in mice with

a high level of K14-Noggin as
judged by real-time quantitative

PCR. The eyelid opening process is not delayed
in our K14-Noggin

mice, but the eyelid opening is smaller. Recently, BMP
pathway

was proposed to be involved in the timing of eyelid opening (Sharov et
al., 2003). On a K5-Noggin background, the eyelid opening was delayed by

20
days, and the suppression by Noggin on eyelid apoptosis and

differentiation was
proposed to be a possible mechanism. Also,

keratin 14 is reported to be partially
replaced by keratin 15.

In our K14-Noggin mice, the level of keratin 14
expression is

not diminished in the eye suture in comparison to the back skin.

However, no abnormalities of the adult eyelid were described

in the K5-Noggin
mice.

151
2.4.4 BMP Regulates the Size of External Genitalia and Integument
Differentiation
External genitalia form from the genital tubercle. The genital

tubercle
differentiates into the penis in males or clitoris

in females (Murakami, et al., 1986;
Hildebrand, et al., 1995). The formation of the genital tubercle, its

outgrowth and
differentiation into either penis or clitoris

is the result of epithelial-mesenchymal
interaction (Kurzrock et al., 1999). In both

males and females, external genital
outgrowth is accomplished

by the formation of the prepuce. WNT, SHH, and
FGF signaling

were shown to be involved in genital morphogenesis (Yamaguchi
et al., 1999; Haraguchi et al., 2000; Haraguchi et al., 2001). Thus,

external
genitalia are another example of ectodermal organs

regulated by a similar set of
morphogenesis-related signaling

molecules. Here we show BMPs, in particular
BMP4, to be expressed at the

tip of the genital tubercle where growth may be
regulated. K14-Noggin

mice of both genders show excessive outgrowth of the
external

genitalia, which is especially apparent in the postnatal period.

We
suggest that BMP signaling regulates the growth of penile

and clitoral tissues in
mice, and that ectopic Noggin disrupts

the balanced growth of these structures
and results in their

hypertrophy. In mice there are hairy spines, which are
epithelial appendages

of the glans penis with possible mechano-sensory function.
Their

formation starts around P10 and it was shown to be androgen-dependent,

because it is primarily retarded in androgen-insensitive mice (Murakami et al.,
1987). Here we showed that BMPs are expressed within the epithelium

of
developing hairy spines and control their differentiation.

Excessive Noggin in the

152
basal layer of the preputial lamellae

inhibits hairy spine maturation, but not the
number of hairy

spines. Therefore, BMP signaling is important for the
differentiation

of hairy spines, but not for their induction and periodic
arrangement.

2.4.5 BMP and Human Diseases
The BMP pathway is of fundamental importance for early stages

of development
(Hogan et al., 1996). Therefore, all mutations of BMPs are likely

to be lethal.
Noggin is a direct antagonist of BMPs. Human loss-of-function

mutations in the
Noggin gene (NOG) were reported. Affected people

have fusion of the joints
(proximal symphalangism, SYM1) or

multiple-synostoses syndrome (SYNS1)
and conductive deafness,

because of stapes ankylosis (Brown et al., 2002; Brown
et al., 2003). To date, no known gain-of-function

NOG gene mutations are known.
However, if they exist, theoretically

these mutations should result in phenotypes
similar to that

of a hypothetical BMP-knockout human. We have searched for
human

congenital anomalies associated with a locus at 17q22, because

human
NOG is mapped to this area of chromosome 17 (Valenzuela et al., 1995). We
found

that several phenotypical features of people with chromosome

17q trisomy
syndrome [mental retardation (MCA/MR) syndrome]

resemble those found in
K14-Noggin mice (Bridge et al., 1985; Sarri et al., 1997). These include:

polydactyly of the hands and feet, syndactyly of the fingers

and toes; hirsutism, a
widows peak (low, v-shaped hair

growth near the top of the human forehead),

153
low posterior hairline,

and external genital abnormalities including a bifid
scrotum

and penile chordae. These abnormalities parallel the paw abnormalities,

hypertrichosis, and external genital abnormalities seen in K14-Noggin

mice. We
speculate that a higher dosage of Noggin, resulting

from an additional NOG gene
allele in people with 17q trisomy

is partially responsible for the above-mentioned
abnormalities.

2.4.6 Morpho-Regulation: Variations or Pathology?
Although some phenotypic features are pathological and result

in loss-of-function
(absence or aberrant claws, replacement

of eccrine glands in paw and Meibomian
glands in eyelids with

hair), many changes are relatively mild and mostly
regulatory

in nature. These changes seem to be quantitative (eg, an increase

in
pelage hair density), qualitative (eg, reduction of the size

of footpads), or
functional (eg, shorter telogen). Although

all these changes are still considered
abnormal because they

indeed significantly deviate from the normal average
phenotypes,

it may be worthwhile to contemplate the borders between pathology

and phenotypic variations. The concept of morpho-regulation implies that
morphogenetic

processes can be modulated by morphological regulators that

lead
to changes of morphological phenotypes in development and

in evolution
(Slavkin et al., 1998). Because the levels of morpho-regulators can

be adjusted
physiologically, they provide means for modulating

the morphology of organs
without drastic changes. Whereas Edelman

concentrated on cell adhesion

154
molecules as morpho-regulators,

here we develop this concept further to major
morphogenesis

signaling pathways (eg, BMP, Wnt, Shh, FGF pathways) and their

modulation by physiological antagonists. Using the pliable BMP

pathway as an
example, this genetically engineered mouse illustrates

the morpho-regulatory
hypothesis vividly. In the context of evolution, the term phenotypic plasticity

is
used to describe the ability of a phenotype to shift quantitatively (Sara et al.,
2001). At the level of species, it may be based on the selection from

a spectrum of
phenotypic variations based on environmental changes.

Examples are seen in the
different densities and length of hairs

observed in mountain cats, dogs, oxen, and
so forth, from temperate

or extremely cold areas found in arctic or high mountain
regions (West et al., 2002), or the shift of finch beak shapes in accord to climate
changes

in Galapagos islands (Kawano et al., 2003). Variations in the number and
size of

integumentary appendages can be used to generate a spectrum

of variable
animal phenotypes that may work as substrates for

selection and become
advantageous when environments change.

However, when these morphological
or structural variations impede

normal functions, they will be considered
pathological.

The recognition that accumulation of mild mutations or variations

can result in the formation of a new trait or new species is

not new, but candidate
molecular pathways responsible for

these variations are mostly unknown. Here
we show Noggin/BMP

antagonism may serve this mechanism. Further study on a
more

quantitative and more regulatory level is needed to develop

this concept
further. In this case, integument appendages constitute

an ideal model because
their changes are usually nonlethal (eg,

unlike many changes affecting heart or
lung) and are easier

to be quantified (Widelitz et al., 1999).

This study shows how

155
simple tuning up and down of the key molecular

pathways activity, such as the
BMP, may regulate the formative

process of ectodermal organs. In the era of
tissue engineering,

one may want to modulate the number, size, or the
differentiation

status of some ectodermal organs in humans or animals for various

medical, agricultural, and industrial reasons. Tissue engineers

will have to learn
how to accomplish the subtle balance of activities

for the major signaling
pathways. The newly made transgenic

mouse can be a useful animal model and
tissue source for these

analyses and evaluations.

156
Chapter 2: Morphoregulation of teeth.
Modulating the number, size, shape and differentiation by tuning Bmp
activity

Summary
During development and evolution, the morphology of ectodermal organs can be
modulated so that an organism can adapt to different environments. We have
proposed that morphoregulation can be achieved by simply tilting the balance of
molecular activity. We test the principles by analyzing the effects of partial
downregulation of Bmp signaling in oral and dental epithelia of the keratin
14-Noggin transgenic mouse. We observed a wide spectrum of tooth phenotypes.
The dental formula changed from 1.0.0.3/1.0.0.3 to 1.0.0.2(1)/1.0.0.0. All
mandibular and M3 maxillary molars were selectively lost because of the
developmental block at the early bud stage. First and second maxillary molars
were reduced in size, exhibited altered crown patterns, and failed to form
multiple roots. In these mice, incisors were not transformed into molars.
Histogenesis and differentiation of ameloblasts and odontoblasts in molars and
incisors were abnormal. Lack of enamel caused misocclusion of incisors, leading
to deformation and enlargement in size. Therefore, subtle differences in the level,
distribution, and timing of signaling molecules can have major morphoregulatory
consequences. Modulation of Bmp signaling exemplifies morphoregulation
hypothesis: simple alteration of key signaling pathways can be used to transform
a prototypical conical-shaped tooth into one with complex morphology. The

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involvement of related pathways and the implication of morphoregulation in
tooth evolution are discussed.

3.1 Introduction
The formation of ectodermal organs depends on a series of
epithelialmesenchymal interactions mediated by signaling pathways. Some
components of these pathways are shared as evidenced by ectodermal dysplasia
syndromes in which hair, teeth, sweat glands, or sometime lungs, all become
defective by the mutation of a single gene (Slavkin et al. 1998; Pispa and Thesleff
2003; Ohazama and Sharpe 2004). We have suggested earlier that different
ectodermal organs are variations sharing a common theme (Chuong 1998). The
sharing of major signaling pathways (e.g., Shh, Bmp, Fgf, Notch, Wnt pathways,
etc.) among the organogenesis of hairs, feathers, teeth, mammary glands, etc.,
attests to the concept of these common themes. Yet how the variations are
generated and regulated are mostly unknown. We surmise the variation is based
on autonomous regional specificity (e.g., Hox codes) and nonautonomous
morphoregulators (e.g., activity of secreted signaling molecules). Identification of
the molecular basis of these variations is ongoing and in this study we focus on
the role of morphoregulation. In contrast to proposing novel molecular pathways,
the concept of morphoregulation postulates that diverse organ phenotypes, in
development or evolution, can be achieved through physiological modulations of
existing morphogenesis-related pathways (Edelman 1992). Although originally
proposed for adhesion molecules as mediators, the concept of morphoregulation
later expanded to cover signaling molecules that work upstream of adhesion

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molecules (Plikus et al. 2004). Direct ablation of a fundamental signaling
pathway is likely to be lethal. However, nature has devised a strategy using a
series of antagonists expressed in a temporalspatial specific way to fine tune
pathway activity and to generate a spectrum of moderate modifications of organ
morphology. We have recently used keratin (K14)-Noggin mice to demonstrate
the basics of this concept (Plikus et al. 2004). We showed that modulation of
Bmp activity indeed leads to various morphoregulation changes in multiple
ectodermal organs including increased numbers of hair filaments, reduced size of
claws, enlarged size of external genitals, and the conversion of sweat glands in
foot pads and Meimobian glands in eyelids into hairs.

To further analyze the principles of ectodermal organ morphoregulation at
different hierarchical morphogenetic levels, we chose one organ to analyze the
consequences of tuning down, but not shutting off, Bmp pathway activity at
different stages of organogenesis. Teeth were chosen for the following reasons:
(i) They show hierarchical levels of morphological complexity resulting from
successive stages of morphogenesis (Thesleff and Mikkola 2002; Tucker and
Sharpe 2004). As teeth in different parts of the oral cavity develop with different
temporal schedules, there is a good chance that different teeth will be affected by
decreased Bmps activity at different developmental stages. (ii) In mammals, there
are regionally specific tooth phenotypes (Sharpe 1995). It has been reported that
the mouse incisor can be re-specified to become a molar in explant cultures
(Tucker et al. 1998) and it would be very interesting to test if this phenomenon
happens in vivo in our K14-Noggin mice. (iii) Teeth have an excellent fossil

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record, and it now is possible to study the roles of genes in development
experimentally, and then match the resulting patterns to dental diversity caused
by natural selection by studying fossils and extant animals. Teeth fulfill all these
criteria more than hairs, feathers, and glands (Yu et al. 2002; Wu et al. 2004), and
therefore have begun to be analyzed from this perspective (Kangas et al. 2004).
(iv) Structural defects in teeth occur often in various human dental diseases. The
versatile phenotype of teeth in K14-Noggin mice may be useful as an
experimental model for pathological studies.

During vertebrate evolution, teeth have been lost and gained. As reptiles evolved
into birds and mammals, teeth met very different fates. Although most Mesozoic
birds had teeth (Hou et al. 2003, 2004), teeth were lost in the Cenozoic era. In the
mammalian lineage, the generally conical-shaped reptilian teeth became more
complex, and mammalian dentition underwent remarkable morphological and
functional diversification, showing great variations of number, size, and shape
(Line 2003). In general, four classes of teeth are arranged from the distal to the
proximal snout, incisors, canines, premolars, and molars, with the number
expressed in sequence as the dental formula. The primitive placental mammals
have a dental formula of 3.1.4.3./3.1.4.3. Some rodents, mice, for example, have
the dental formula, 1.0.0.3/1.0.0.3, with only one incisor, no canines, no
premolars, and three molars. Homologous teeth can greatly range in size among
different mammalian species. Incisors in many species such as humans are
relatively small. In contrast, incisors in rodents can reach a large size in relative
proportions (Tummers and Thesleff 2003). The shape of teeth within the same

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class can also vary. Molars and premolars generally have a more complicated
crown pattern than canines and incisors, and are well understood from a
functional perspective and are usually related to specific molar functions such as
shearing, crushing, and grinding of food (Hiiemae 2000). Although there are
functional reasons for the expansion of mammalian teeth diversity, the gain, and
loss of teeth, the ways to achieve that remain mostly unknown. This is
particularly intriguing as quite different morphologies can occur in closely related
clades (e.g., the aforementioned mouse and vole), whereas similar changes (e.g.,
loss of teeth in baleen whales, pangolins, and anteaters) occur independently in
different clades. On a larger scale, these phenomena suggest that the tooth
morphogenetic pathway is in a quasi-stable equilibrium in individual species and
that it is sensitive to molecular tuning, resulting in phenotypic plasticity.

Many growth factors (Fgf8, Egf, Tgfb1, Bmp2, Bmp4, etc.) and transcription
factors (Msx1, Msx2, Pax9, Lef1, etc.) were shown to play key roles during
various phases of odontogenesis (reviewed in Thesleff and Sharpe 1997;
Scarel-Caminaga et al. 2002; Thesleff and Mikkola 2002; Tucker and Sharpe
2004). It is believed that diversification of the dentition is achieved through the
modulation of activities and timing of these pathways, as well as differences in
requirements by various tooth primordia (Thesleff and Sharpe 1997; Jernvall and
Thesleff 2000). Among the multiple signaling pathways known to regulate tooth
development, the Bmp pathway stands out for its importance (Bei and Maas
1998, 2000; Reddi 1998; Zhang et al. 2000; Miyazono et al. 2001). Various Bmps
are expressed throughout odontogenesis. They exhibit a complex expression

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pattern (Aberg et al. 1997; Yamashiro et al. 2003). Overall, Bmp2, Bmp4, and
Bmp7 expression patterns largely overlap. Bmp3 and Bmp5 show a rather
distinct and restricted distribution in tooth compartments. Most interestingly,
inhibition of Bmps signaling with noggin changes tooth identity from incisor to
molar. Prior to odontogenesis embryonic day 9 (E9)(E10) Bmp4 inhibits Barx1
expression in the presumptive incisor mesenchyme. In contrast, Barx1 expression
is stimulated by Fgf8 in the presumptive molar mesenchyme. If Bmps activity is
experimentally downregulated by exposing the E9E10 mandibular arch to
exogenous noggin, incisors transform into molars (Tucker et al. 1998). This
compelling experiment has prompted us to look further into how normal Bmp4
signaling may regulate epithelialmesenchymal interactions and specify the fate
and shape of tooth primordia. In this study we use noggin as a tool to tune down
the activity of the Bmp pathway. By overexpressing noggin in the oral and dental
epithelium, we expected to disturb the otherwise balanced activity of Bmp
pathway in odontogenesis. As noggin is secreted, mis-expressed noggin should
have a nonautonomous effect on both dental epithelium and dental mesenchyme,
blocking Bmp signaling in epithelialmesenchymal interactions. Indeed we found
a broad spectrum of changes in different aspects of odontogenesis. We describe
these changes in detail and discuss them in the context of tooth development,
diseases, and in the broader context of the morphological evolution of ectodermal
organs.

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Figure 3.1 Production of K14-Noggin mouse.
K14 Noggin construct used to generate transgenic mouse. The size of inset used
and restriction enzyme are indicated (A). Appearance of control C57BL/6J (B)
and mutant K14-Noggin 2.5-month-old mice (C). Examples of K14
immunostaining is shown in newborn molars (D, F) and incisors (E, G) which are
strongly K14 positive. Section plane: sagittal (AH). Scale bars: 100 µm (E); 50
µm (D); 10 µm (F, G).

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Figure 3.1: Continued

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3.2 Materials and Methods
3.2.1 Production and genotyping of transgenic mice
Mice were generated in the Norris Cancer Center transgenic mouse facility at the
University of Southern California as previously described (Figure 3.1, AC
adopted from Plikus et al. 2004). All phenotypic features of K14-Noggin mice
showed high penetrance. All animals were treated under humane conditions
following protocols approved by the University of Southern California IACUC.

3.2.2 Histological and immunochemical staining
Tissues were collected and fixed in 4% paraformaldehyde, dehydrated, embedded
in paraffin, and sectioned at 56 µm. When necessary, specimens were
additionally decalcified after fixation. Standard H&E staining was performed for
basic histological analysis. Immunostaining was performed using the Ventana
DiscoveryTM automated immunostaining module (Ventana Medical Systems,
Tucson, AZ, USA). Primary antibodies used were rabbit anti-PCNA (1:500,
Santa Cruz Biotechnology, Santa Cruz, CA, USA) and rabbit anti-K14 (1:400,
Berkeley Antibody Company, Richmond, CA, USA). The DAB (Ventana
Medical Systems) or HistoStain (Zymed Laboratories, San Francisco, CA, USA)
detection kits were used for color development.

3.2.3 In situ hybridization
Mouse tissues from various ages were used for in situ hybridization. Tissues for
in situ samples were fixed and dehydrated in DEPC-treated solutions according to
a standard protocol. To detect mRNA expression, the tissue was hybridized with

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the appropriate digoxigenin-labeled probe. Signals were detected using an
anti-digoxigenin antibody coupled to alkaline phosphatase. Some tissue samples
were processed using the Ventana DiscoveryTM automated in situ hybridization
instrument (Ventana Medical Systems).

3.2.4 Scanning electron microscopy (SEM) analysis
Tissues were prepared according to the standard SEM protocol. Briefly, samples
were fixed in 2.5% glutaraldehyde in 0.1 m sodium cacodylate, dehydrated, and
critical point dried from ethanol. Samples were coated with gold in a sputter coat
chamber. SEM was performed in the Doheny Eye Institute Core Facility at the
University of Southern California.

3.3.5 Ground sections analysis
After fixation and dehydration, tissues were embedded in Eponate 12 (Ted Pella
Inc., Redding, CA, USA). Curing was done at 60°C for 48 h. The
resin-embedded specimens were ground all the way to the middle of the teeth.
The surface was then polished using a fine sharpening stone.

3.3 Results
3.3.1 K14 promoter activities in oral cavity
Basal K5 and K14 are first detected in E9.5 in mice. β-galactosidase driven by
human KRT5 promoter was seen in the first branchial arch as early as E10.5
(Byrne et al. 1994). We examined K14 immunohistochemistry in the oral
epithelium at E13 (not shown). In the incisor bud, K14 was expressed in the

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proximo-labial part of the bud, but was absent from the distallingual part.
Similarly in the E13 molar, K14 was predominantly expressed in the proximal
part of the bud. As tooth development progressed, levels of K14 expression in
molars increased but remained low in the E15.5 incisor (not shown). Expression
of K14 stayed high in the oral epithelium. At postnatal day 1 (P1), K14
expression was high throughout the oral epithelium, the ameloblast cell layer, and
the stratum intermedium both in molars (Figure 3.1D and 3.1F) and incisors
(Figure 3.1E and 3.1G). K14 in preameloblast and ameloblast was also detected
by immunochemistry (Tabata et al. 1996). Expression levels of endogenous K14
in the K14-Noggin mice were similar to those of the WT teeth (not shown).

3.3.2 The number of K14-Noggin molars is reduced
Both genotypically and phenotypically two groups of K14-Noggin mice could be
identified: low-transgenic (TG) copy number and high-TG copy number animals
(Plikus et al. 2004). High-TG copy number K14-Noggin mice exhibited overall
more dramatic pathological changes in various skin appendages (Figure 3.1C).
Therefore, in our present study we analyzed the dental phenotype in high-TG
copy number (HCN) K14-Noggin mice only.

Mandibular molars. The most dramatic finding is that K14-Noggin mice lack
all mandibular molars. K14-Noggin mice show consistent changes of the dental
formula to 1.0.0.2(1)/1.0.0.0 (Figure 3.2AH). All three pairs of mandibular
molars were absent in all (n=24) animals studied. On histology, thickened and

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Figure 3.2 Early arrest of K14-Noggin molar development.
(AD) WT mice have three maxillary (A) and three mandibular molars (C). Most
of the K14-Noggin mice have only two or rarely one maxillary molar (B). None
of the K14-Noggin mice have mandibular molars (D). (EH) WT mice have three
maxillary (E) and three mandibular molars (G) with a well-developed crown and
roots. K14-Noggin mice have a reduced number of maxillary molars (F) and no
mandibular molars. Instead, there is thickened oral epithelium, often resembling
residual molar lamina (H). (IL) Developing molars in the E13 WT (I, K) and
K14-Noggin (J, L) mouse embryos. In WT mice, tooth buds have subjacent
mesenchymal condensations (I, K). In K14-Noggin mice, the maxillary molar
bud has a distinct mesenchymal condensation (outlined with green dotted line).
However, the mandibular molar bud (L) lacks a mesenchymal condensation.
Epithelium is outlined with red. (MP) E15 molars in the WT (M, O) and
K14-Noggin (N, P) mouse embryo. Unlike maxillary molars (N), K14-Noggin
mandibular molars fail to develop further (P). Epithelium resembles initial tooth
lamina (red). There is no mesenchymal condensation beneath it (green arrow). (Q,
R) Bmp4 expression in the mesenchyme of the developing K14-Noggin teeth at
E13 (Q) and E15 (R). (S, T) Msx1 expression pattern during early odontogenesis
in K14-Noggin mice. In WT mice at E13, Msx1 is distinctly expressed in the
dental mesenchyme, but not in the epithelium (S, inset). In E13 K14-Noggin mice,
Msx1 is expressed in the dental mesenchyme of the maxillary molars (S). Msx1
expression is seen in the mandible, but not directly underneath the oral epithelium
(S). At E15, when K14-Noggin mandibular molars show developmental arrest,
Msx1 is expressed in the maxillary dental mesenchyme, as well as in the

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Figure 3.2: Continued
mesenchyme of the arrested mandibular early bud (T). (U, V) Absence of the
neuro-specific differentiation of the dental mesenchyme in both maxillary and
developmentally arrested mandibular tooth buds (V), as judged by the Tubb3
expression. Fibers of the trigeminal nerve are strongly positive for Tubb3 (see
inset on U). Section plane: sagittal (EH, J, L, N, P, QV), frontal (I, K M, O).
Scale bars: 1 mm (AH), 100 µm (IV). de, dental epithelium; dm, dental
mesenchyme.

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Figure 3.2: Continued

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Figure 3.2: Continued

171
invaginated oral epithelium was present in the adult transgenic animals (Figure
3.2H). There were no indications of teeth or teeth-like structures.

Maxillary molars. The number of maxillary molars is also reduced in
K14-Noggin mice. The third maxillary molars (M3) were absent with very high
penetrance (>95%). In two out of 24 mice, the second molars (M2) were missing,
one on the left side and the other on the right side (Figure 3.2B). The first molars
(M1) were always present in all (n=24) animals included in this study.

Developmental changes. To further determine the stage where the
developmental block occurs, we studied morphogenesis of both maxillary and
mandibular M1 molars of the K14-Noggin mice. At E13.5 M1 molars of the WT
mice were in the bud stage (Scarel-Caminaga et al. 2002). Oral epithelium
formed bud-like downgrowth surrounded by mesenchymal condensation (Figure
3.2I and 3.2K; Maas and Bei 1997). K14-Noggin maxillary M1 molars showed
bud stage morphology (Figure 3.2J). In contrast, mandibular molars were blocked
at the early bud stage showing only a thickened dental epithelium, which did not
seem to invaginate. There were only slight increases in the cellular density of the
underlying mesenchyme (Figure 3.2L).

In WT mice, molars entered cap stage at E14.5 and further progressed to the bell
stage at E16.5 (Figure 3.2M and 3.2O). At E15, K14-Noggin maxillary M1
molars appeared to be at the early cap stage. Epithelial tooth buds seemed to fold,
but did not form a distinctive cap-like structure surrounding the mesenchymal

172
papilla (Figure 3.2N). Transition from the bud to cap stage was delayed in
maxillary molars. Mandibular molars did not progress in their development any
further. They remained early bud-like in appearance, exhibiting thickened dental
epithelium with little or no mesenchymal condensation beneath it (Figure 3.2P).
To ensure that the effect seen is because of noggin activity, the expression of
Bmp4 was confirmed by in situ hybridization (Figure 3.2Q and 3.2R). No
differences were found between the WT and K14-Noggin mice. In a similar
manner, the expression of Msx1 was examined as it has been suggested to
mediate epithelialmesenchymal interactions during tooth induction (Chen et al.
1996). We found Msx1 to be expressed in the mesenchymal component of the
E13 WT tooth buds (Figure 3.2S, inset). In E13 K14-Noggin embryos, Msx1
expression was seen in both maxillary and mandibular incisor buds (not shown)
and throughout the maxillary molar mesenchyme. In the mandible, Msx1 showed
mesenchymal expression, yet not directly under the dental epithelium (Figure
3.2S). By E15, Msx1 was strongly expressed in the dental mesenchyme of
maxillary molars and only weakly in the mesenchyme adjacent to the mandibular
dental placode (Figure 3.2T). The results suggest that Msx1 expression during the
early development of K14-Noggin molars remains largely normal.

Another marker examined was tubulin, β3 (Tubb3). In Msx1-deficient mice
dental mesenchyme differentiated abnormally and was reported to express the
neuronal marker Tubb3 (Han et al. 2003). In E15 K14-Noggin mice, neither of
the dental mesenchyme from the maxillary molars and mandibular laminae
expressed Tubb3 (Figure 3.2U and 3.2V). As a control, the trigeminal nerve was

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strongly Tubb3 positive (Figure 3.2U, inset). These results suggest that the
developmental inhibition caused by noggin could be an event occurring later than
Msx1-related blockage. Alternatively a non-Msx1-mediated pathway is used.

We then examined whether rates and distribution of apoptosis are altered in these
mutants. E15 tooth buds from both WT and K14-Noggin were TUNEL negative,
suggesting the absence of apoptosis (not shown). At the same time, other areas of
the embryos were TUNEL positive (not shown). No TUNEL-positive cells were
found within the K14-Noggin mandibular lamina structure at E15 (not shown).

3.3.3 Defects in K14-Noggin maxillary molars include abnormal crown/root
patterning and enamel/dentine differentiation

Crown size/pattern. Compared with the WT, K14-Noggin maxillary molars
were significantly smaller (Figure 3.3J vs. 3.3K). There was a reduction in the
size of the crown base and low proliferation rates. Miniaturization was obvious
both at P21, when K14-Noggin molars started to erupt (Figure 3.3A vs. 3.3B),
and in adulthood. Overall morphology of the crown was changed as well (Figure
3.3G vs. 3.3H). Widths of both K14-Noggin M1 and M2 molars at the neck level
were about 53% less than that in WT. Reduction in the crown size was generally
associated with low rates of proliferation. Although normally the growing area of
the cervical loop is highly Pcna positive (Figure 3.7G), it was virtually devoid of
proliferating cells in P1 K14-Noggin molars (Figure 3.7H).

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Figure 3.3 Growth, eruption, and patterning and differentiation defects in
K14-Noggin molars.
(A, B) Comparative morphology of the maxillary molars in the P21 WT (A) and
K14-Noggin mice (B). WT molars have a well-developed crown. M1 and M2
WT molars have well-developed multiple roots. Unlike WT teeth, maxillary M1
and M2 molars in K14-Noggin mice are smaller, have no clear crown to root
separation, and do not form multiple roots. Postnatal development of the
K14-Noggin maxillary molars is largely retarded. (CF) Delayed eruption of
molars in K14-Noggin mice. Although P21 M1 and M2 WT molars have fully
erupted (C, D), both maxillary molars in P21 K14-Noggin mice remain at the
early stages of eruption, with the crown mostly seated deep within the alveolae
(E, F). (GI) Abnormal crown pattering in K14-Noggin maxillary molars. P21
WT maxillary molars have a well-defined cusp pattern, with seven cusps in M1
and five cusps in the M2 molar (G). P21 K14-Noggin M1 and M2 maxillary
molars have a severely abnormal crown pattern with small cusp-like prominences
(H). Position and number of the intercusps are inconsistent in P21 M2
K14-Noggin molars (inverted view, I). (J, K) K14-Noggin molars have multiple
root defects. WT molars have multiple well-developed roots (J). K14-Noggin
molars have either two short, misconfigured roots (M1), or only one, very short
root (M2), and multiple, irregular, grape-like growths on their surface (K). In WT
molars, root length dominates over crown length (J). In K14-Noggin molars, the
crown/root to multiple root ratio is changed and the crown/root length dominates
over multiple root length (K). (LS) Comparative morphology of P8 (LO), P14
(PS) WT, and

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Figure 3.3: Continued
K14-Noggin maxillary molars. Both in WT and K14-Noggin molars HERS is
clearly seen at P8 (N, O). At P14 HERS in K14-Noggin molars has largely
normal morphology (S vs. R). Although roots start to form at or before P8, furcae
does not form until much later in K14-Noggin molars. (TW) Formation of small
furcae in P28 K14-Noggin molars. In M1 K14-Noggin molars furcae is first seen
at P28 (V). Furcae is delineated by a layer of disorganized, mineralized dentin
(W). Periodontal apparatus forms with periodontal ligament connected to the
cementum (V, inset). Not all K14-Noggin molars form furcae and multiple roots
(U vs. T). (X, Y) Crown surface defect in K14-Noggin molars. Unlike in WT
molars, the crown surface of the K14-Noggin adult molars is uneven and highly
eroded (X). On ground sections, WT molars show a distinct layer of enamel (Y,
green arrows), whereas no enamel is seen in K14-Noggin molars (Y). Section
plane: sagittal (LW, Y). Scale bars: 1 mm (A, B, G, H, J, K); 0.5 mm (I);
200 µm (L, M, P, Q, T, U); 100 µm (V, X, Y), 50 µm (N, O, R, S, W). AB,
aleveolar bone; am, ameloblasts; bv, blood vessels; c, cementum; d, dentin; dpm,
dental papillae mesenchyme; e, enamel; f, furcae; HERS, Hertwig's epithelial root
sheath; od, odontoblasts; p, pulp; pdl, periodontal ligament.

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Figure 3.3: Continued

177
Figure 3.3: Continued

178
Figure 3.3: Continued

179
Other areas of the crown, such as ameloblasts and stratum intermedium of the
intercusps also showed low proliferation rates in K14-Noggin teeth (Figure 3.7L
vs. 3.7K). On the other hand, elevated proliferation was seen in the epithelium
and preodontoblasts of the presumptive cusps in P1 K14-Noggin molars (Figure
3.7J vs. 3.7I). Proliferation rates in the epidermis and developing hair follicles of
the P1 WT and K14-Noggin mice were virtually the same (Figure 3.7B vs. 3.7A).
We found the crown pattern in maxillary molars to be altered. Normal crowns
form discrete cusp and intercusp regions. In adult mice, the maxillary M1 and M2
molar have seven and five cusps, respectively (Figure 3.3G). K14-Noggin molars
appeared to develop blunt cusp morphology (Figure 3.3B). However, progressive
deterioration prevented accurate assessment of crown patterning in adult mutants
(Figure 3.2B). The crown pattern was studied at P21 instead. At P21,
K14-Noggin molars were just starting to erupt and were not affected by
mechanical wearing and caries (Figure 3.3E and 3.3F). On SEM, individual cusps
were clearly visible in P21 WT molars (Figure 3.3G). In contrast, no distinct
cusps could be identified in K14-Noggin mice. Instead, small, partially fused
prominences formed (Figure 3.3H). We also compared several P21 M2 molars
and found the number and position of intercusps to be inconsistent even among
molars from the same animal (Figure 3.3I). The eruption of K14-Noggin molars
was significantly delayed. At P21, both K14-Noggin maxillary molars were just
starting to erupt (Figure 3.3E and 3.3F). At the same time, all three maxillary
molars were fully erupted in WT mice (Figure 3.3C and 3.3D).

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Root size/pattern. At P8, WT maxillary molars showed formation and
downward migration of the Hertwig's epithelial root sheath (HERS) at the apical
end of the molars at which time the shape of the roots starts to appear (Figure
3.3L and 3.3N). In contrast, P8 K14-Noggin maxillary molars although having
formed the HERS structure, showed no evidence of root formation (Figure 3.3M
and 3.3O). By P14, the roots of both M1 and M2 WT maxillary molars were
formed and were covered by a layer of cementum. The periodontal ligament
(PDL) connecting the cementum and the alveolar bone was clearly seen (Figure
3.3P and 3.3R). At P14, K14-Noggin molars progressed little and the only sign of
root formation was the presence of HERS (Figure 3.3Q and 3.3S). However, the
forming roots of the K14-Noggin molars had largely reduced rates of
proliferation, both within the HERS and dental mesenchyme (Figure 3.7R vs.
3.7Q). At the same time, rates of proliferation were comparable in the oral
epithelium adjacent to molars (Figure 3.7N vs. 3.7M). There was still no
evidence of furcae formation that signifies the splitting of one root into multiple
roots (Figure 3.7Q). At P21, WT maxillary M1 molars had well-formed roots,
with the root length nearly equal to the length of the crown. M2 molars had
formed distinct roots, but they were shorter than the crown.

M3 molars had initiated root development (Figure 3.3A). Both maxillary M1 and
M2 K14-Noggin molars were lacking any signs of root bifurcation (Figure 3.3B).
At P28, the WT molars were in the process of erupting with the enamel exposed
to the oral cavity. There was a clear cemento-enamel junction (CEJ) demarcating
the ending of the crown and the beginning of the roots, which had completed

181
their development. The periodontal apparatus was in place (Figure 3.3T). In the
K14-Noggin molars, there was no CEJ and no clear separation between the crown
and the roots (Figure 3.3U); however, there was a layer of cementum and PDL
present. Some M1 K14-Noggin molars formed a very small furcae, which
indicates the formation of the different roots (Figure 3.3V and 3.3W). Hence, we
called this structure a crown/root (Figure 3.3K).

Although all WT maxillary molars had three distinct roots (Figure 3.3J), mutant
M1 maxillary molars showed some signs of root bifurcation (Figure 3.3K),
whereas M2 molars did not form multiple roots and, in essence, consisted of
crown/roots. In addition, irregular, bud-like outgrowths were observed over the
surface of mutant molars (Figure 3.3K). Crown/root to multiple root proportions
were reversed in K14-Noggin molars (Figure 3.3J vs. 3.3K). In adult M1 and M2
WT molars, the crown to multiple root ratio was 0.53 and 0.85, respectively. This
ratio became 1.8 in M1 adult K14-Noggin molars (Table 3.1).

Enamel/dentin differentiation. WT molars are white, smooth, and shiny.
K14-Noggin molars were dull, gray, and showed numerous, widespread
caries-like lesions, suggesting a poorly mineralized enamel structure (Figure 3.2B
vs. 3.2A). On SEM, adult mutant molars showed multiple macro- and
microscopic sites of dental decay (Figure 3.3X). Ground sections showed they
did not have an enamel layer (Figure 3.3Y).

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At E18, M1 WT molars entered the late bell stage and soon started ameloblast
and odontoblast differentiation. K14-Noggin maxillary M1 molars appeared to be
still at the early bell stage at P1. The epithelialmesenchymal interface of the P1
WT molars was highly structured. From the epithelial side there was a layer of
polarized ameloblasts interfacing at the basal end by the stratum intermedium.
From the mesenchymal side there was a layer of dentin-producing polarized
odontoblasts (Figure 3.4A). In contrast, the epithelialmesenchymal interface of
the K14-Noggin molars at P1 did not show a similar specialization. There were
no polarized ameloblasts but rather multiple layers of disorganized
preameloblasts. There were no signs of odontoblasts on the periphery of the
dental papillae. There were no enamel or dentin depositions (Figure 3.4B). To
further determine the developmental status of the K14-Noggin molars, we studied
the expression of differentiation markers Amelx (amelogenin) and dentin
sialophosphoprotein (Dspp). Expression of Amelx in ameloblasts marks the onset
of the secretory stage and was present at P1 in WT teeth (Figure 3.4C). In
contrast, P1 molars in K14-Noggin mice did not express Amelx (Figure 3.4D).
Dspp expression starts at the late bell (differentiation) stage and was present in P1
WT teeth in the preodontoblasts/odontoblasts. Consistent with previous reports,
some Dspp expression was seen in the preameloblasts (Figure 3.4, E and G;
Begue-Kirn et al. 1998). In K14-Noggin P1 molars Dspp expression was largely
absent (Figure 3.4F and 3.4H). However, dentin-like material was seen in
K14-Noggin molars starting from P8 (Figure 3.3I). These deposits of dentin-like
material were associated with the late expression of the odontoblasts-specific
Dspp markers, as seen at P14 (Figure 3.4J vs. 3.4I).

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Figure 3.4 Epithelio-mesenchymal defect in K14-Noggin molars.
(AD) Crown surface defect in K14-Noggin molars. Unlike in WT molars (A),
the crown surface of the K14-Noggin adult molars is uneven and highly eroded
(B). On ground sections, WT molars show a distinct layer of enamel (C, green
arrows), whereas no enamel is seen in K14-Noggin molars (D). (A, B)
Epitheliummesenchyme interface of the P1 WT (A) and K14-Noggin (B) M1
molars. Unlike WT, K14-Noggin molars stay largely undifferentiated. (CJ)
Absence of ameloblast-specific and delayed expression of odontoblast-specific
markers. In P1 WT teeth, Amelx is strongly expressed in the ameloblasts,
especially in the cusp (C). Dspp has strong expression in the preodontoblasts and
odontoblasts, as well as in the preameloblasts of the intercusp (E, G). In P1
K14-Noggin molars, both Amelx (D) and Dspp (F, H) are largely absent.
K14-Noggin molars gain Dspp expression later. Strong Dspp expression is seen
at P14 (I vs. J). Scale bars: 500 µm (DF); 200 µm (I, J); 100 µm (AC, G, H).
Section plane: sagittal (AJ). am, ameloblasts; de, dental epithelium; dp, dental
papilla; ide, inner dental epithelium; od, odontoblasts; pd, predentine; si, stratum
intermedium.

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Figure 3.4: Continued

185
Dspp expression starts at the late bell (differentiation) stage and was present in P1
WT teeth in the preodontoblasts/odontoblasts. Consistent with previous reports,
some Dspp expression was seen in the preameloblasts (Figure 3.4, E and G;
Begue-Kirn et al. 1998). In K14-Noggin P1 molars Dspp expression was largely
absent (Figure 3.4F and 3.4H). However, dentin-like material was seen in
K14-Noggin molars starting from P8 (Figure 3.3I). These deposits of dentin-like
material were associated with the late expression of the odontoblasts-specific
Dspp markers, as seen at P14 (Figure 3.4J vs. 3.4I).

3.3.4 Defects in K14-Noggin incisors happen predominantly during late
morphogenetic events
WT mice have one pair of incisors in the upper and lower jaw with a shiny,
semi-transparent, yellowish surface (Figure 3.5A and 3.5I). In K14-Noggin mice
all incisors were always present, but were thick, wide, blunt ended, and
misaligned with a dull white surface (Figure 3.5B and 3.5JL). They deteriorated
and developed marked unilateral erosions because of constant toothwear between
upper and lower pairs. These changes started early and became more severe. The
surface of K14-Noggin incisors was rough and defective (Figure 3.5, E and 3.5F
vs. 3.5C and 3.5D). It showed both macroscopic signs of deterioration in the form
of deep, parallel ridges (Figure 3.5E), and microscopic irregularities in the form
of multiple bud-like formations (Figure 3.5F). On the ground sections, the labial
side of WT incisors displayed a clear, thick layer of enamel (Figure 3.5G). In
contrast, K14-Noggin incisors do not have any enamel (Figure 3.5H).

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Figure 3.5 Growth abnormalities of the K14-Noggin incisors caused by the loss
of enamel.
(A, B) Comparative gross morphology of the mandibular incisors in the adult WT
(A) and K14-Noggin mice (B). K14-Noggin incisors are thick, wide, blunt ended,
and misaligned. (CF) On SEM, the surface of the K14-Noggin incisors is rough
and defective (E, F). It shows both macroscopic signs of deterioration in the form
of deep, parallel ridges (E) and microscopic irregularities in the form of multiple
bud-like formations (F). The surface of WT incisors is smooth (C, D). (G, H) On
ground sections, WT incisors display a clear, thick layer of enamel (G). In
contrast, K14-Noggin incisors do not have any enamel layer present (H). (IL)
Progressive changes of the incisors in K14-Noggin mice. Unlike WT incisors (I),
K14-Noggin incisors are a dull white and deteriorate because of constant rubbing
against each other (JL). These changes start early in life and with age become
more severe. The bottom incisors grew very long and became needle sharp (L).
Section plane: sagittal (G, H). Scale bars: 100 µm (C, E, G, H); 20 µm (D, F).

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Figure 3.5: Continued

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Early developmental stages of the K14-Noggin incisors seemed to be unaffected.
Similar to the WT incisors, at E13, K14-Noggin incisors were at the bud stage
and by E15 further progressed into the cap stage (not shown). Developmental
defects became apparent at later differentiation and secretion stages. At P1, WT
incisors showed clear specialization of the dental epithelium into the ameloblasts
and stratum intermedium. Peripheral dental papilla differentiated into
odontoblasts. Sandwiched layers of enamel and dentin were seen in between
ameloblasts and odontoblasts (Figure 3.6A). From the labial side of the growing
end, actively proliferating cells were confined to the small area of the cervical
loop (Figure 3.7E). Differentiation of the P1 K14-Noggin incisors was delayed.
Retarded differentiation was associated with a markedly expanded zone of
proliferation on the labial side of the cervical loop (Figure 3.7F vs. 3.7E).
K14-Noggin incisors formed irregular layers of preameloblasts and
preodontoblasts (Figure 3.6B). Preameloblasts lacked clear parallel cell
alignment, did not express Amelx (Figure 3.6E and 3.6F vs. 3.6C and 3.6D), and
did not deposit enamel. Preodontoblasts looked poorly differentiated and at that
stage did not express Dspp (Figure 3.6I and 3.6J vs. 3.6G and 3.6H), but seemed
to deposit predentin-like material (Figure 3.6B). The Epitheliamesenchymal
border had an irregular, wavy appearance throughout the K14-Noggin incisors
(Figure 3.6B, arrowheads). It is notable, however, that K14-Noggin P1 incisors
were generally more differentiated than P1 molars (Figure 3.4F vs. 3.6B).

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Figure 3.6 Differentiation defect in K14-Noggin incisors.
(A, B) Epitheliummesenchyme interface of the P1 WT (A) and K14-Noggin (B)
M1 incisors from the labial side. Unlike WT incisors, K14-Noggin incisors
remain less differentiated. They do not form a distinct layer of polarized
ameloblasts and there is no dentin or enamel deposition.
Epitheliummesenchyme interface is uneven and wavy. (CJ) Absence of the
tooth-specific differentiation markers in the early postnatal (P1) K14-Noggin
incisors. Amelx is strongly expressed in the ameloblasts of the WT incisors (C,
D). In WT P1 molars Dspp has distinct expression (G). Dspp is expressed in the
preodontoblasts and odontoblasts, as well as in the preameloblasts of the
proximal labial side (H). In K14-Noggin P1 incisors, both Amelx (E, F) and Dspp
(I, J) are largely absent. (K, L) Epitheliummesenchyme interface of the P14 WT
(K) and K14-Noggin (L) M1 incisors from the labial side. K14-Noggin incisors
remain poorly differentiated. Epitheliummesenchyme interface is uneven and
wavy. Section plane: sagittal (AL). Scale bars: 100 µm (AL). am, ameloblasts;
dp, dermal papilla; od, odontoblasts; pam, preameloblasts; pod, preodontoblasts;
pd, predentine; si, stratum intermedium.

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Figure 3.6: Continued

191
Figure 3.7 Proliferation defect in K14-Noggin teeth.
(A, B) Similar rates of proliferation in the epidermis and hair follicles of P1 WT
(A) and K14-Noggin (B) mice. (CF) Proliferation pattern in P1 WT (C, E) and
K14-Noggin (D, F) incisors. Proliferation rates are comparable on the lingual side
of the cervical loop of both WT (C) and K14-Noggin (D) incisors. However, the
proliferation zone on the labial side of the K14-Noggin cervical loop is
significantly expanded distally (F vs. E). (GL) Proliferation pattern in P1 WT (G,
I, K) and K14-Noggin (H, J, L) molars. K14-Noggin molars show greatly
reduced rates of proliferation, especially in the cervical loop (H vs. G) and
intercusp area (L vs. K). (MR) Reduced and nonlocalized proliferation activity
in P14 K14-Noggin molars. WT molars show a localized zone of proliferation at
the tip of the cusps (O) and extensive proliferation activity within HERS and the
surrounding dental mesenchyme (Q). In contrast, K14-Noggin molars have
reduced, de-centralized proliferation activity within the dental epithelium of the
crown (P), largely reduced proliferation in HERS, and virtually no proliferation
in the surrounding dental mesenchyme (R). Contrarily, the oral epithelium,
adjacent to teeth, shows comparable proliferation activity both in WT (M) and
K14-Noggin mice (N). Section plane: sagittal (AR). Scale bars: 50 µm (AR).
am, ameloblasts; dp, dental papilla; ide, inner dental epithelium; od, odontoblasts;
pod, preodontoblasts; si, stratum intermedium, sr, stellate reticulum.

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Figure 3.7: Continued

193
Figure 3.7: Continued

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3.4 Discussion
3.4.1 Bmp signaling is required for the growth of molars and crown
morphogenesis
Bmp signaling is critical for molar progression through the early bud stage. At
E13, the K14-Noggin mandibular molar lamina does not progress to form a
distinct bud, and only a dental lamina-like epithelial thickening remains. While at
E13 there is some increase in the dental mesenchymal density beneath the lamina,
it disappeared by E15. Despite the morphological abnormalities in dental
mesenchyme, we are surprised to find that K14-Noggin mice have largely normal
Msx1 expression. We also found mesenchymal Bmp4 expression in both
maxillary and mandibular K14-Noggin molar regions at E13. During early molar
development Bmp4 and Msx1 were shown to cooperate closely (Chen et al.,
1996). It is generally believed that epithelial Bmp4 signals through Msx1 in the
mesenchyme to induce its mesenchymal expression of Bmp4, as supported by the
fact that Msx1-deficient mice express epithelial but not mesenchymal Bmp4.
Tooth development in these mutants does not progress past the bud stage (Bei
and Maas, 1998; Chen et al., 1996). We suggest that K14-Noggin mandibular
molars are developmentally blocked during mesenchymal Bmp4 signaling back
to the dental epithelium, when the mis-expressed noggin abolishes this signaling
cascade.

During normal odontogenesis, molecular signals from primary enamel knots
determine the size of the tooth crown by coordinating dental epithelia
proliferation and folding, as well as the position of secondary enamel knots. The

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location of secondary enamel knots defines normal crown patterning in
multi-cusp molars (Jernvall et al., 1994). Crown defects in K14-Noggin mice are
similar to, but more severe than in K14-Follistatin mice (Wang et al., 2004a). In
follistatin-deficient mice, lack of activin / Bmps antagonist, results in reduced
inner dental epithelium proliferation, irregular folding, and shallow, un-polarized
cusps (Wang et al., 2004a). Bmps might also control the distance between
adjacent secondary enamel knots, thus regulating the positioning of the cusps
(Jernvall and Thesleff, 2000). Here we observe that K14-Noggin produces a
disturbed distribution of growth centers in incisors and molars. In the
K14-Noggin molar crown, proliferation is abnormally low in the epithelium and
mesenchyme within the cervical loop, leading to miniaturized maxillary molars.
K14-Noggin molars develop very small cusp-like prominences that are partially
fused without clear intercusps. Paradoxically, the K14-Noggin incisor produces
an abnormal but more diffuse distribution of proliferation. Therefore there are
region specific responses to the same stimuli (see below).

3.4.2 Bmp signaling is required for roots patterning and growth
A recent report showed Bmp2, Bmp4, and Msx2 in the HERS cells of P10 mice
by in situ hybridization (Yamamoto, 2004). It appears that proliferation of HERS
in the K14-Noggin mice is reduced resulting in delayed root formation. Even
more pronounced is the defect in root patterning. The formation of the furcae that
serves to delineate the different forming roots is largely delayed in M1 molars
and never occurs in M2 molars. M1 K14-Noggin molars showing a small furcae,
form what appears to be two small rudimentary roots, while their wild type

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counterparts form three distinct roots. The combined root defect and absence of a
well delineated cemento-enamel junction result in an indistinct crown-root
border.

Growth factors such as Tgfb1 and its receptors (Gao et al., 1998; 1999), Bmp2,
Bmp3, and Bmp7 (Thomadakis et al., 1999) have been found in cementoblasts,
periodontal ligament, and alveolar bone. However, until now no direct functional
role in root formation has been demonstrated for these factors. The involvement
of Dlx3 in root development is supported by the phenotype expressed by patients
with tricho-dento-osseous (TDO) syndrome. The K14-Noggin dental phenotype
has some resemblance to TDO in that the maxillary molars have an altered
crown-to-root ratio and very short malformed roots (taurodontism). A nonsense
mutation in the DLX3 gene was identified in a family with TDO syndrome (Price
et al, 1998). Amongst defects in hair, bone, and enamel, patients with TDO
syndrome also present root defects. It is interesting that Bmp2 has been shown to
regulate transcription of Dlx3 in keratinocytes (Park and Morasso, 2002). These
results are consistent with the notion that Bmps are required for root formation.

3.4.3 Bmp signaling is essential for the histogenesis of enamel and the dentin
layer.
Our results suggest that Bmp signaling controls the maturation of the
epithelial-mesenchymal interfaces in developing teeth. It appears that, after the
morphogenesis phase, Bmps are critical for instructing the inner dental
epithelium to stratify and form ameloblasts during the differentiation phase. In

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K14-Noggin mice, the epithelial-mesenchymal border is disorganized and the
inner dental epithelium fails to polarize and stratify. Ectopic noggin prevents
normal differentiation of the ameloblast lineage. The pre-dentin layer contains
several secreted growth factors that stimulate the differentiation of ameloblasts
and secretion of amelogenin. Bmps, such as Bmp2, are known to be transcribed in
odontoblasts (Aberg et al., 1997), and can induce ameloblast differentiation (Coin
et al., 1999b). Odontoblast-derived Bmps are believed to stimulate ameloblast
differentiation through the induction of p21 and Ambn (ameloblastin; Wang et
al., 2004b). Additionally, Bmps are present in tumors that form enamel, dentin,
cementum, or bone (Gao et al., 1997). Here we provide supportive in vivo
evidence that Bmps are indeed required for ameloblast differentiation. Follistatin
was shown to act as the main inhibitor of the Bmps-driven ameloblast
differentiation in the lingual dental epithelium of mice incisors (Wang et al.,
2004b). Asymmetric expression of follistatin accounts for the presence of the
enamel on the labial side only. Overexpression of noggin throughout the dental
epithelium of the K14-Noggin incisors, wipes out asymmetry of Bmp pathway
activity, and results in incisors free of enamel on both the lingual and labial sides.
Signaling along the epithelial-mesenchymal interface is reciprocal. Ameloblasts
in turn secrete growth factors that stimulate osteogenesis and cementogenesis in
the adjacent tissues. Enamel extracts were shown to contain an osteo-inductive
ability, which is reduced when pre-incubated with Bmp2 / Bmp4 antibody or
noggin (Iwata et al., 2002). K14-Noggin teeth fail to develop odontoblasts in a
timely fashion. Although layers of dentin-like material eventually form, they are
deposited irregularly and are functionally defective. The dentin phenotype could

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be interpreted either as a direct effect of noggin or indirectly via the lack of
proper differentiation of the ameloblast layer. Whatever the mechanism, the
results show the essential importance of Bmps in the differentiation of the enamel
and dentine layers.

3.4.4 Differential effects of Bmp signaling along the dental axis
We are somewhat disappointed to find the absence of the incisor to molar
conversion in K14-Noggin mice (Tucker et al., 1998). It is possible that this is the
result of timing and strength of the KRT14 promoter activity. The KRT14
promoter may not be active or active enough in the oral epithelium over the
presumptive incisor region at E9-E10 (Byrne et al., 1994). In fact, unlike in
molars, the KRT14 promoter has low activity in incisors even at the cap stage
(E15). Incisors show a dramatic increase in KRT14 activity at later developmental
stages (P1). Indeed, late developmental defects dominate in K14-Noggin incisors.
Still we observed region specific responses to noggin by teeth in different molar
forming regions. These differences cannot be simply explained by the temporal
difference of tooth development, and may imply some fundamental differences in
their development or evolutionary basis. More work is obviously required, but
these results are consistent with the hypothesis that mandibular teeth have
different signaling pathway requirements during their development (Thomas et
al., 1997). By tuning the levels of Bmps activity, our results suggest that
maxillary and mandibular molars have different levels of requirements for Bmp
signaling and that mandibular molars show higher dependency on Bmp signaling
in the early bud stage. Maxillary molars, albeit abnormal, proceeded through the

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early bud stage with reduced level of Bmps activity, while mandibular molar
primordia did not form at all. However, maxillary molars still require some Bmps
activity, as complete abolishment of Bmp signaling in Bmpr1a-deficient mice
blocks the development of all teeth (Andl T et al., 2004).

Among the maxillary molars, we observed inhibition of odontogenesis more on
the distal, but less on the mesial end of the dental axis. The third maxillary molar
almost always fail to develop and the second maxillary molars are sometimes
missing. We propose three possible mechanisms for this phenomenon. First, the
third molar develops later than the first two (M1 goes through bud stage at E13,
M2 - at E16, and M3 only upon birth), and it is possible that by the newborn
stage, KRT14 promoter activity in the dental epithelia has risen high enough to
completely block development of the tooth bud. Our K14 immunostaining data
support this hypothesis. Second, early Bmp signaling deficiency may reduce the
size of the committed molar field and there is simply not enough cellular mass
left to form the third molar (Wang et al., 2004a). The more provocative third
possibility is that either epithelial or mesenchymal cells forming the maxillary
M3 molars are intrinsically different from those forming M1 and M2 molars, and
may have similarly high Bmp signaling dependencies like the mandibular molars.
Indirect evidence suggests that neural crest cells along the jaw axis may be
intrinsically different (Ruch, 1995; Sharpe, 1995). Neural crest cells originate at
different time points from topologically different areas. Maxillary molar crest
cells appear to derive from the anterior midbrain at the 5-somite stage and
posterior midbrain at the 6-somite stage (Imai et al., 1996; Köntges, and

200
Lumsden, 1996). Mandibular molar crest cells are predominantly derived from
the posterior midbrain and partially from the anterior hindbrain at the 5/6-somite
stage. We speculate that distal maxillary mesenchyme may behave similar to the
mandibular dental mesenchyme. Indeed different dental regions have different
competence based on their distinct intra-cellular molecular compositions. Barx1
is expressed in molar mesenchyme and determines molar phenotype. Isl1
(homeobox gene; synonym: Islet 1) is expressed in incisor epithelium and
determines incisor phenotype (Mitsiadis et al., 2003). For the maxilla/mandible, it
is shown that in response to the epithelial Fgf8, maxillary mesenchyme response
with Dlx2 expression only, and mandibular mesenchyme response with both Dlx2
and Dlx5 expression. This was used as evidence for the existence of internal
differences of maxillary and mandibular mesenchyme, probably due to
differences in their origin from the neural crest (Ferguson et al, 2000).

3.4.5 Comparison with other mouse tooth mutants and human dental
diseases
Genetically engineered mice involving the Bmp pathway have been generated.
Mice with a deletion of Bmpr1a in their epithelium driven by the KRT14
promoter lack all teeth upon birth. Development of both molars and incisors is
arrested at the bud stage (Andl et al., 2004). Since teeth are arrested at early
stages of development, there is no opportunity to evaluate the role(s) of Bmp
signaling in later events of histogenesis and differentiation. On the other hand, no
dental phenotype was documented for Bmp5 (Kingsley et al., 1992) and Bmp7
(Dudley et al., 1997) loss-of-function mutations. This could be due to the

201
redundancy of Bmp ligands; multiple, if not all Bmps have to be inactivated to
disturb normal odontogenesis. Mice deficient for noggin or Chrd (another Bmps
antagonist; synonym: chordin) also do not demonstrate significant dental
phenotypes (Stottmann et al., 2001). Under physiological conditions noggin is
apparently not expressed during early dental development, however, follistatin
and perhaps ectodin (ectodermal Bmp inhibitor; aka: Sostdc1, sclerostin domain
containing 1) antagonize Bmps signaling in developing teeth (Laurikkala et al.,
2003). In the teeth of other genetically engineered or mutated mice, there are
several interesting findings related to our observation (Figure 3.8C). All teeth in
Lef1- (Kratochwil, 1996), Pitx2- (Szeto, 1999), Pax9- (Peters et al., 1998),
Msx1/Msx2- (Bei and Maas, 1998), Gli2/Gli3- (Hardcastle et al., 1998) deficient
mice are also completely absent, implying a block in the earlier inductive stage.
These molecules would not be ideal molecular tools for fine tuning tooth
morphology in the context of morphoregulation. There are also selective losses of
tooth types. In activin mutant mice, incisors and mandibular molars fail to
develop beyond the bud stage, while their maxillary molars are unaffected
(Ferguson et al., 1998). Mice over-expressing follistatin (Tgfb inhibitor) under
the KRT14 promoter lack both maxillary and mandibular third molars, have a
disturbed cusp pattern, and show premature wearing of enamel (Wang et al.,
2004a). Dlx1/Dlx2-deficient mice do not form maxillary molars, but their incisors
and mandibular molars are normal. It was shown that the mutated
ectomesenchyme underlying the maxillary molar regions loses its odontogenic
potential and forms chondrocytes instead (Qiu et al., 1997; Thomas et al., 1997).

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Figure 3.8 Summary of the multiple dental defects caused by the disruption of the
Bmp pathway in the oral epithelium.
(A) Summary diagram of the tooth developmental events affected by the reduced
strength of Bmp signaling in K14-Noggin mice. (B) Comparison of the dental
formulas between WT, K14-Noggin, and other known mutant mice with
reduction in teeth number.

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These results further corroborate the hypothesis that there are both qualitative and
quantitative differences among different teeth.

In various dental diseases, humans also suffer from, malformation abnormal
differentiation, and loss of teeth. Humans have the dental formula of
2.1.2.3/2.1.2.3. Agenesis of one or several teeth (hypodontia) is a common
congenital defect (Vastardis, 2000). Less prevalent is oligodontia, when six or
more permanent teeth are missing. Mutations in transcription factor genes MSX1
and PAX9 were found responsible for the nonsyndromic forms of oligodontia
(Vastardis et al. 1996; Stockton et al. 2000; Nieminen et al. 2001; Lammi et al.
2003). Syndromic oligodontias are often associated with multi-organ syndromes,
and like hypodontia, they are believed to have heterogeneous and mostly
non-established genetic backgrounds. In addition to tooth agenesis, congenital
dental pathology also enlists defects in dentin differentiation (e.g. dentinogenesis
imperfecta type II; Zhang et al., 2001), enamel differentiation (e.g. amelogenesis
imperfecta; Lagerstrom et al., 1991), root formation (e.g. hypoplasia of teeth
roots; Lind, 1972), teeth eruption (Stoelinga et al, 1976), etc. There are also
non-genetic human dental diseases involving defects in the enamel, dentine, and
cementum. The diverse dental phenotypes in K14-Noggin mice may become a
useful experimental model to study the progression, consequences, side effects,
and management of these defects in vivo.

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3.4.6 Evo-Devo Implications of teeth and other ectodermal organ
Among ectodermal organs, teeth are the most common elements found in the
fossil record. A large body of literature on vertebrate paleontology describes
evolutionary changes in tooth morphology (Crompton, 1995; Hiiemae, 2000).
While these changes are well-understood from a functional perspective, the
developmental mechanisms behind these changes are mostly unknown. The
evolutionary origin of molar shape and dentition patterns was previously
summarized (Crompton, 1971). In archetypical reptiles, the teeth have one single
elevation (or a prototypic cusp) which is triangular in shape with a cylindrical
base. There is not much regional diversity in the oral cavity. Most Mesozoic birds
have teeth along their entire jaw (e.g., Archeopteryx; Feduccia, 2001), or only in
the distal beak (e.g., Longirostravis; Hou et al., 2004). The morphology of their
teeth is of the prototypical conical shape without any signs of regional diversity.
Modern and Cenozoic birds as well as some Mesozoic birds (e.g. Confucisornis)
lack teeth completely (Hou et al., 2003). As mammal-like reptiles evolved, their
dentitions became more complex (Crompton, 1995, Hiiemae, 2000). From the
prototypic dental formula of 3.1.4.3./3.1.4.3., loss of teeth is a common theme in
mammalian evolution, while addition is rare. In some cases, teeth are lost
completely (e.g., baleen whales), or all teeth of one jaw are lost (e.g., upper jaw
of the spermwhales). More frequently, a whole class of teeth is missing, such as
the loss of the upper incisors in deer (Cervidae), or of the premolars in mice.
Changes in crown patterning are associated with different modes of food
processing. Examples were described above in the mice and voles (Tummers and
Thesleff, 2003). Loss of regional specificities can result in similarity of all teeth

205
along the tooth row (e.g., armadillos, Dasypodidae; dolphins, Delphinidae). In
different animals, there are different proportions and arrangements of hard tissues
(enamel, dentin, and cementum), because of their different material properties
(Martin et al., 2003). In grass-eating herbivores (e.g., horses, antelope), enamel
forms high-crowned molars, with valleys filled in with dentin and cementum for
grinding. On the other hand, enamel is present in scanty quantities on elephant
tusks, which wear off soon, and teeth are reduced to dentin pegs. With new
understanding in tooth formation, we can learn more on how these diversities can
be generated by variations in the induction (gain or loss of number),
morphogenesis (patterning), or differentiation (ratio of hard tissues) stages. This
Bmp pathway study provides some clues that can be verified in extant animals
(Figure 3.8A, 3.8B).

Some efforts have been made to understand the Evo-Devo of teeth. Tooth loss
has happened frequently and independently during evolution. Developmentally,
how was this achieved? Are there multiple ways to loose teeth? To see if tooth
can be rescued in avians, Kollar recombined chicken oral mucosa and mouse
dental mesenchyme (Kollar and Fisher, 1980). Chicken dental lamina can
progress to the bud stage when Bmps/Fgfs are added to the organ culture.
Chicken oral epithelium is competent to form tooth like appendage follicles when
it is adjacent to feather forming mesenchyme (Chen et al., 2000) or mouse
cephalic crest cells (Mitsiadis et al., 2003). Another tooth region is the diastema
between incisors and molars in mice. It seems to be achieved through the
localized sequestration of Shh by Gas1 in the diastema mesenchyme, causing

206
ablation of the Shh signaling necessary for tooth development (Cobourne et al.,
2004). Thus loss of teeth can be achieved through the modulation of the activity
of several different signaling pathways, and noggin-induced tooth loss may also
have been used.

Another molecular pathway that behaves like a morphoregulator is the
ectodysplastin (Eda) pathway (Srivastava et al., 1997; Tucker et al., 2004). It is
interesting that molar phenotype of the K14-Noggin mice is somewhat similar to
that of Tabby mutant mice. Tabby mice carry a spontaneous mutation in the Eda
(anhidrotic ectodermal dysplasia) gene and show multiple developmental defects
in the ectodermally derived organs, such as hairs, sweat glands, and teeth. Tabby
molars are small. Some cusps are missing; others are either fused or too closely
positioned to each other. In Tabby molars, the primary enamel knot is small,
which results in the smaller crown base and ultimately in the fusion of the
secondary enamel knots. Additionally, growth of the cervical loop epithelium in
Tabby molars is slow. This results in slow formation of the tooth crown base
(Pispa et al., 1999; Jernvall and Thesleff, 2000). With the increased level of Eda
in the tooth forming region, Eda activity imposes a moderate yet persistent effect
on the number of teeth, cusps, and crown complexity (Kangas et al., 2004).
Together with its roles on other ectodermal appendages (Pispa and Thesleff,
2003), the Eda pathway can be considered as another morpho-regulatory
pathway.

207
Our understanding in the Evo-Devo of ectodermal organs has just begun
(Chuong, 1998; Pispa and Thesleff, 2003). Interpreting the transgenic
morphology in the context of the fossil record and analyzing the molecular basis
of tooth diversity in extant mammals beyond the mouse will add to our
appreciation of the fundamental mechanisms of epithelial appendage
morphogenesis. Here we demonstrated that the Bmp pathway is one of the major
morpho-regulatory pathways. We showed that tooth characteristics can be
sculptured by tuning Bmps activity at different stages of tooth morphogenesis.
The repetitive use of the Bmp pathway is consistent with the concept of co-opting
an existing molecular pathway for evolutionary novelty in a different hierarchical
level to reach new complexity (True and Carroll, 2002; Prum and Dyck, 2003).
The study of the morpho-regulatory principles in ectodermal organs will help our
long term objectives of how to guide epithelial stem cells to form organs we
desire.

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Chapter 3: BMP activity in spreading of hair cycle regenerative waves

Summary
The hair cycle can by regulated by molecular oscillations within the follicle
(autonomously) or by dynamic changes in the extra-follicular environment
(non-autonomously). While human hair follicles largely cycle autonomously, the
cycling of mouse hair follicles is affected by the status of extra-follicular
environment and neighboring hair follicles. Coordination of hair cycle among
large groups of hair follicles results in wave-like progression of anagen initiation
and formation of hair cycle domains. Observations of the dynamic hair cycle
domains on the skin of wild type and Msx2
-/-
mice show that each hair cycle
domain is made of an initiation site, a propagating wave, and boundaries.
Analyses of patterns of wave propagation and boundary formation led us to
identify four new functional phases of the hair cycle. Telogen was divided into
refractory and competent phases on the basis of the ability of hair follicles to
re-enter anagen upon stimulation. Anagen was divided into propagating and
active phases on the basis of the ability of hair follicles to propagate anagen
signals onto competent telogen hair follicles. We have conclusively identified
skin-wide changes in Bmp signaling activity as the major regulatory factor in the
new patterned hair cycling mechanism. Newly identified expression dynamics of
Bmp2/4 and noggin within multiple extra- and intra-follicular sites collectively
define Bmp signaling output onto the large populations of hair follicles. High
Bmp signaling levels keep telogen hair follicles in a refractory state. The
subsequent downfall in Bmp signaling strength coincides with the acquisition of

209
telogen competence. A low level of Bmp signaling is maintained during early,
propagating anagen. Anagen propagates among competent telogen hair follicles
and stops at the interface with the refractory telogen follicles, where a visible
anagen-telogen border forms. Consequently, a sharp uprise in Bmp2/4 expression
during anagen V-VI boosts Bmp signaling levels and terminates the propagation
properties of anagen. Loss of these propagation properties allows for the
maintainance of borders between domains. The spatial distribution of large skin
domains in different functional stages leads to the formation of observed
macroscopic patterns, and constitutes the basis of patterned hair growth
phenomenon. Our work further demonstrates the non-genetic, non-inherited, but
rather dynamic nature of this pattern formation process.

4.1 Introduction
Previously we have described Msx2
-/-
mice that exhibit the cyclic alopecia
phenotype (Ma L et al., 2003). Cyclic alopecia uncovers an intriguing
phenomenon of patterned hair growth by allowing us to visualize macroscopic
distribution patterns of anagen and telogen hair follicles that form on the surface
of the skin. These patterns are complex, resemble geometric shapes and change
over time. Many other mutant mice with the cyclic alopecia phenotype have been
reported to have patterned hair growth as well (Koch et al., 1998; Militzer, 2001;
Chidgey et. al, 2001; Suzuki et al., 2003; Uyttendaele et al., 2004; Mammucari
et al., 2005). However, despite multiple accounts of patterned hair growth almost
nothing is known about the underlying patterning mechanism and the molecular
signals involved. Basic observations of patterned hair growth suggest some type

210
of inter-follicular hair cycle coordination mechanism. This mechanism should be
distinct from the two other known modes of hair cycle regulation: autonomous
and synchronous. Autonomous regulation characterizes lack of hair cycle
harmonization between neighboring hair follicles. Under this mode of regulation,
each follicle appears to cycle autonomously, as seen in human scalp and guinea
pig pelage (Van Steensel et al., 2000). In synchronous growth mode, the hair
cycles of all hair follicles are regularly brought in-sync by a systemic, typically
endocrine, signal. Estrogens and prolactin have a profound systemic influence on
the hair cycle. In many animals, including mice, estrogens, such as 17ß-estradiol
(E2), inhibit anagen initiation (Fraser and Nay, 1953) by targeting follicular
estrogen receptors (Ohnemus et al., 2005). Withdrawal of estrogens by
gonadectomy or administration of an estrogen receptor

antagonist (ICI 182.780)
stimulates premature anagen (Chanda et al., 2000; Oh and Smart, 1996; Smart et
al., 1999). Prolactin is another potent systemic modulator of hair growth (Pearson
et al., 1996; Pearson et al., 1999; Nixon et al., 2002). Commonly, in seasonal
animals, the increase in pituitary prolactin during spring induces new anagen
(Dicks, 1994). Simultaneously, prolactin can be produced locally and it can
regulate the hair cycle in a non-systemic way (Foitzik et al., 2003; Craven et al.,
2001).

The phenomenon of patterned hair growth has been well appreciated in classical
studies on rats, mice and other mammals (Durward and Rundall, 1949; Mottram,
1945). Early observations in rats have described hair growth patterns as
successive waves of anagen periodically spreading from the ventral side of the

211
body to the dorsal side, over the trunk (Durward and Rundall, 1949; Mottram,
1945) and progressively decreasing in width with age (Durward and Rundall,
1949; Butcher, 1936). The periodic nature of these hair-growth waves was
thought to be inherent and was largely attributed to some genetic, yet unidentified
mechanism (Durward and Rundall, 1949; Whiteley, 1958; Ebling and Johnson,
1959). Conversely, it was demonstrated that the so-called inherent rhythms of
hair growth can be modulated by systemic factors, such as steroid hormones
(Butcher, 1936; Ebling and Johnson, 1961; Ebling and Hervey, 1964; Ebling and
Hale, 1966). Experimentation with steroid hormones led to the identification of
the important phenomenon of telogen refractivity (Ebling and Johnson, 1961;
Johnson, 1958a; Johnson, 1958b). Telogen refractivity was defined by the simple
fact that a systemic influence can induce new anagen only at a particular time,
and that there is a time period following anagen during which the systemic
stimulus is unable to exert an effect (Durward and Rundall, 1949; Ebling and
Johnson, 1961). At that time it was not possible to conclusively identify and
prove the cellular and molecular bases underlying telogen refractivity. Early work
left unanswered the question of whether loss of telogen refractivity involves a
lowered concentration of an inhibitor or an increased concentration of an
accelerator (Ebling and Johnson, 1961). Untill now our understanding of this
phenomenon and the phenomenon of patterned hair growth remains rudimentary.
The mounting morphological evidence for patterned hair growth requires a new
explanation of hair cycle control in large populations of follicles.

212
4.2 Materials and Methods
4.2.1 Animals
C75BL/6J, CD1, C3H/HeJ, SCID and predominantly black Crl:ZUC-Lepr
fa
rats
were used in this study. Msx2
-/-
(Msx2
tm1Rilm
/Msx2
tm1Rilm
), KRT14-NOG,
Bmp4-lacZ, NOG-lacZ, 52bpMsx2-hsplacZ and TOPGAL transgenic mice were
also used.

4.2.2 Hair cycle observation
Progression of the hair growth patterns was monitored in mice and in rats for
various intervals of time, up to 1 year. Changes were documented every 48 hours
and pictures were taken. The fur of all wild type mice was periodically clipped,
as it was growing. Hair clipping was selected over plucking and shaving since
plucking disrupts the normal hair growth cycle and artificially synchronizes hair
growth, while shaving can result in abrasions and wounding that can potentially
interfere with normal hair growth (Chase, 1954; Paus, 1998).

4.2.3 Animal procedures
All procedures were performed on anaesthetized animals (Ketamine
HCl:Xylazyne mixture was used). For skin transplantation, surgical procedures
were performed when both donor and recipient skins were in early telogen. This
was done to ensure that wounded skin is healed by the beginning of the next
anagen phase and that the impact of wound healing on the hair cycle is minimal
(skin wounding in late telogen can induce new anagen around the wound; Plikus
and Chuong, personal observations). SCID mice were used as recipients of skin

213
transplants to assure lack of immunological rejection. After surgery, mice were
monitored for a prolonged period of time. Changes in hair growth patterns were
documented and pictures were taken.

4.2.4 Histology and detection of molecular expressions
Tissues were collected, fixed and processed for histology as described previously
(Plikus et. al, 2004). Whole mount in situ hybridization procedures on thin slices
of adult mouse skin were performed

using the InsituPro automated in situ
detection module (Intavis

AG, Koeln, Germany). Analysis was performed
according to the

standard whole mount in situ protocol (Jiang et al., 1999). The
position of sebaceous glands on the surface of the skin was visualized using a
whole mount staining procedure based on fat staining with Sudan IV dye (Park et
al., 2001). This dye stained sebaceous glands red. Whole mount
beta-galactosidase detection was performed on adult skin from Bmp4-lacZ,
NOG-lacZ, 52bpMsx2-hsplacZ and TOPGAL mice. Staining was performed
according to previously described protocol (Brugger et al., 2004).

4.3 Results
4.3.1 Large groups of pelage hair follicles form macroscopic hair cycle
domains
To identify hair cycle dynamics in large groups of hair follicles in the living
mouse we used two non-invasive assays that can reveal recognizable traits of the
specific hair cycle stages. The first is based on pigmentation. In pigmented mice,
such as inbred C57BL/6J mice, melanin production is restricted to the hair

214
follicles and not to inter-follicular skin. Melanogenesis starts at anagen IIIa,
becomes prominent in anagen IIIb, and continues until catagen (so-called:
anagen-coupled melanogenesis; Muller-Rover et. al, 2001; Figure 4.1B, 4.1D).
Therefore, hair follicles during most of anagen appear grey / black, while hair
follicles in telogen have no pigment and the skin becomes pink. When hairs are
clipped immediately above the skin surface, differences in pigmentation of the
proximal hair follicles becomes apparent (Slominski and Paus, 1993; Figure
4.4B). The second detection method is based on the recently described "cyclic
alopecia" phenotype (Ma et al., 2003). In Msx2
-/-
mice with cyclic alopecia the
hair filaments dislodge at catagen, making telogen skin regions alopecic until hair
follicles regenerate during the next anagen. Both detection methods reveal
patterns of hair growth, consisting of skin patches with hair follicles in telogen or
various stages of anagen. Because these skin patches differ from one another on
the basis of the hair cycle stages they are termed hair cycle domains. The patterns
of hair cycle domains are not limited to a particular mouse strain. We have
observed these patterns in outbred white CD1, inbred agouti C3H/HeJ and
immunodeficient SCID mice. hair follicles becomes apparent (Slominski and
Paus, 1993; Figure 4.4B). The second detection method is based on the recently
described "cyclic alopecia" phenotype (Ma et al., 2003). In Msx2
-/-
mice with
cyclic alopecia the hair filaments dislodge at catagen, making telogen skin
regions alopecic until hair follicles regenerate during the next anagen. Both
detection methods reveal patterns of hair growth, consisting of skin patches with
hair follicles in telogen or various stages of anagen.

215
Figure 4.1 Hair cycle domain and new spatial-temporal expression profiling of
the hair cycle.
A, B) Spatial distribution of hair cycle stages within the hair cycle domain:
schematic representation (A) and whole mount view (B). The blank arrows point
to the direction of the spreading waves.
C, D) Inverted view of the hair cycle domain. The blank arrow points to the
direction of the spreading wave.
E, F, G) Differential expression of TOPGAL (E), Notch1 (F) and Msx2 (G) in
whole mount preparations of hair cycle domains.
H) Schematic overlay of Bmp4, Bmp2 and Noggin expression levels, and Bmp
activity onto the morphological components of the hair cycle domain and
corresponding functional phases of the hair cycle. Colored solid areas denote
strong expression; colored striped areas indicate loss of expression from some but
not all expression sites.

216
Figure 4.1: Continued

217
Rodents other than mice also show hair cycle domain patterns. For example, after
clipping, the predominantly black Crl:ZUC-Lepr
fa
rats show recognizable
pigmented domains (Zucker rats).

We have analyzed the dynamics of hair cycle domain formation as well as the
structure of these domains on whole mount and histological sections (Figure
4.1A-G). Analysis of the domains across their boundaries allowed us to visualize
the distribution of hair follicles in a continuum of hair cycle stages (Figure
4.1C-D). Each domain can be in one of two alternative states: growth (various
stages of anagen) and quiescence (telogen). During growth three principal
components of the hair cycle domain can be identified: an initiation center, a
spreading wave and a boundary (Figure 4.1A). The group of hair follicles in the
most advanced stage of anagen within the domain represents the "initiation
center". Regions of skin with gradients of hair follicles ranging from anagen V to
anagen I form the spreading wave. The zone between anagen I and telogen hair
follicles forms the "wave front" where telogen hair follicles enter anagen. The
"boundary" of the domain forms when a region of skin in telogen does not
respond to the wave-front signals. This boundary can be recognized because it is
flanked by anagen VI and telogen hair follicles.

4.3.2 Telogen is divided into two functional phases: refractory and
competent
Hair cycle dynamics within domains suggest that the passing of an initiation
signal between neighboring hair follicles plays an important role in the

218
propagation of anagen induction over large distances. For this anagen induction
wave to eventually stop and for a sharp morphological boundary to form there
should be an effective mechanism for either abrupt cessation of initiation signal
production or for the lack of competence of the hair follicles to respond to this
signal. Our data suggest that early telogen hair follicles are indeed refractory to
anagen initiation. On many occasions all hair follicles from the domain in early
telogen are not induced to re-enter anagen by the signals from a new initiation
center adjacent to it. They form a sharp telogen/anagen border with the
surrounding skin whose hair follicles are competent and propagate the induction
signal from the initiation center (Figure 4.2A). Similar phenomena can be
achieved experimentally by offsetting the hair cycle to early telogen within a
group of hair follicles that is otherwise surrounded by hair follicles in late telogen
(Figure 4.2B). We offset the cycle by stimulating extranumerary anagen in hair
follicles that receive cyclosporin A treatment (cyclosporin A is a known anagen
stimulatory agent; Maurer et. al, 1997). Upon completion of cyclosporin
A-induced anagen, we obtain a group of hair follicles whose telogen is ~13 days
younger than that of the surrounding hair follicles (12.7 days is the average
length of anagen for dorsal hair follicles, Figure 4.2E). When a new initiation
center forms, an anagen induction wave starts to propagate throughout the skin by
the end of the second week (counting from the time of the completion of
cyclosporin A-induced anagen) but the offset hair follicles remain in telogen (i.e.
they are refractory; Figure 4.2B, day 15).

219
Figure 4.2 Experimental identification of refractory and competent telogen
phases.
A) Natural refractory telogen. Telogen domain (arrowhead) is refractory to the
anagen stimulatory signal originating from the initiation center (dot) and
spreading throughout competent telogen skin around it (arrow).
B) Experimental induction of refractory telogen achieved by offsetting the hair
cycle within small skin area to early telogen stage.
C) Competence-dependent response of telogen skin to anagen stimulation
established through club hair plucking. Arrowheads point at the site of new hair
growth.
D) Comparison of response time of competent and refractory telogen skin to club
hair plucking stimulation illustrated on 2C.
E) Timing of normal telogen length reveals the presence of an initial refractory
phase, i.e. minimal telogen (~28 days), during which new anagen can not be
initiated.

220
Figure 4.2: Continued

221
To establish the duration of refractory telogen we have performed long term
measurements of hair cycle stages for the dorsal and ventral domains of
C57BL/6J mice (Figure 4.2E). While anagen length in dorsal hair follicles
appears to be tightly regulated (12.7 +/- 0.97 days (n=18)), telogen length shows
great variability (59.6 +/- 18.7 days (n=22)) with a minimal length of 28 days and
a maximal length of 88 days. Data suggest that new anagen can not be initiated
until 28 days pass by, during which time telogen hair follicles remain refractory
to natural anagen stimulatory signals. After that, hair follicles become competent
and can re-enter anagen at any time point. Similar telogen timing dynamics have
been found in the ventral skin. While anagen length is only 7.6 +/- 0.66 days
(n=28) which is about two times shorter than anagen in dorsal hair follicles, it is
also tightly controlled around that value. At the same time telogen length is 40.3
+/- 10.3 days (n=30) and it spans from a minimum of 25 days to a maximum of
78 days. Here again data suggests that the first 25 days represent a refractory
telogen phase.

To further establish the dependence of anagen stimulation from the competence
of the telogen hair follicles we have designed a semi-quantitative club hair
plucking assay. We have plucked either 50 or 200 club hairs and we did so either
during early, refractory or late, competent telogen phases (club hair plucking is
know to stimulate anagen; Collins, 1918; Silver and Chase, 1970). Plucking
resulted in new anagen induction at different time points, depending on the initial
status of the telogen follicles (Figure 4.2C and 4.2D). Competent telogen follicles
respond

222
the earliest (by day 8) to the plucking of 200 club hairs. Similarly, competent
telogen follicles show delayed response to the plucking of 50 club hairs (after 14
days). On the other hand, refractory telogen follicles show a significantly delayed
response to the plucking of both 200 and 50 club hairs (only after 42 days).

The above results indicate that upon completion of anagen, each hair follicle
enters refractory telogen. Refractory telogen is essential for the formation of
domain borders and it probably determines the observed macroscopic hair cycle
domain patterns. Refractory telogen has a fixed length and is followed by a
competent telogen phase of variable length. The transition into competent telogen
is associated with recommencement of the ability of a hair follicle to re-enter
anagen either spontaneously (as an initiation center), as the result of lateral
induction by neighboring anagen hair follicles (as part of the spreading wave), or
in response to extraneous anagen stimulation (such as club hair plucking).

4.3.3 Anagen is divided into two functional phases: propagating and active
The dynamics of anagen propagation suggest that early anagen hair follicles
(anagen I-III) at the tip of the spreading wave consistently and quickly induce
competent telogen hair follicles ahead of the spreading wave into new anagen.
This is how the spreading wave propagates. At the same time, late anagen hair
follicles (anagen VI) most often do not stimulate competent telogen hair follicles.
This phenomenon can be observed when the anagen spreading wave stops and
forms a boundary with the refractory telogen hair follicles. Over the time period
that refractory follicles re-enter the competent phase, simultaneously, early

223
anagen follicles on the other side of the boundry progress into anagen VI. Despite
these changes competent telogen hair follicles remain quiescent. Anagen VI hair
follicles can not stimulate them. This phenomenon is responsible for the
stabilization of domain boundaries. On the basis of this phenomenon we have
divided anagen into two phases: propagating (early, inducing) and active (late,
non-inducing).

4.3.4 Level of Bmp signaling correlates with the maintenance of the
refractory status by telogen hair follicles
Functional differences between refractory and competent telogen phases should
be associated with differential expression of either inhibitor or activator signaling
pathway(s) within and/or around telogen hair follicles. By screening for the
candidate signaling pathways we have identified members of the Bmp signaling
pathway: Bmp4 and Bmp2 show differential expression patterns correlating with
temporal changes in the competence status during telogen. Specifically, we have
identified that refractory telogen is associated with high levels of Bmp2 and
Bmp4 expression in various compartments of telogen follicles and in the
surrounding interfollicular skin. Competent telogen is associated with the loss of
most of these Bmp4 and Bmp2 expression sites. We have identified Bmp4
expression by studying Bmp4-lacZ transgene activity in large whole mount skin
samples containing several hair cycle domains (Figure 4.3H-Q). During
refractory telogen Bmp4-lacZ is abundantly expressed in the dermal papillae and
secondary hair germs of the telogen hair follicle, in the epidermis, dermal
fibroblasts, and arrector pili muscles of the interfollicular skin (Figure 4.3I, 4.3L).

224
In situ expression of Bmp4 and Bmp2 was studied on long thin-slice whole mount
skin samples stretching over several hair cycle domains. Bmp4 in situ mimics
Bmp4-lacZ transgene activity. Bmp2 is abundantly expressed in subcutaneous
adipocytes underlying telogen hair follicles during refractory telogen (arrowhead
on Figure 4.3C and 4.3F'). We have confirmed Bmp2 expression to be in
subcutaneous adipocytes by its overlapping staining with Sudan red marking
fat-containing adipocytes (Figure 4.3G, 4.3G'). Upon transition into late,
competent telogen all of the above Bmp4-lacZ expression sites are lost (Figure
4.3K, 4.3N). Most prominently, Bmp4-lacZ expression disappears from the
dermal papilla. Complete loss of Bmp4-lacZ expression is preceded by a
transition stage with limited Bmp4-lacZ expression patterns in the sebaceous
glands, infundibulum epidermis and secondary hair germs (Figure 4.3J, 4.3M). At
this stage Bmp2 almost completely disappears from the subcutaneous adipose
tissue (arrowheads on Figure 4.3D). We have performed in situ staining on
adjacent long thin-slice whole mount skin samples to identify the level of
temporal/spatial overlap of the above described Bmp4 and Bmp2 expression
dynamics during telogen. Results demonstrate a high level of overlap. Refractory
telogen skin is characterized by strong Bmp4 expression in the dermal papillae
(Figure 4.3R) and simultaneous strong Bmp2 expression in the subcutaneous
adipocytes (Figure 4.3U). Both Bmp4 (Figure 4.3S) and Bmp2 (Figure 4.3V)
expression sites disappear in competent telogen skin.

225
Figure 4.3 Spatial and temporal changes of Bmps expression in the hair cycle
domain.
A-E) Differential in situ expression of Bmp2 in extrafollicular tissue. Positive in
situ signal is pseudocolored in green. Yellow arrows point at Bmp2 expression
during anagen (B, C, D). Red arrowheads point at Bmp2 expression during
refractory telogen (C, E) or lack of Bmp2 expression during competent telogen
(D). The blank arrows point to the direction of the spreading waves; --| signs
indicate position of anagen-telogen border (A, C, D) or competent-refractory
telogen border.
F-G) Extrafollicular Bmp2 expression in subcutaneous adipocytes based on
morphology (F, F'; red arrowheads point at the site of Bmp2 expression) and by
the overlapping staining with Sudan red marking fat-containing adipocytes (G,
G'; yellow arrows point at the overlapping sites of Bmp2 in situ and Sudan red
staining).
H-Q) Differential Bmp4 expression throughout hair cycle stages as demonstrated
by Bmp4-lacZ transgene activity. Loss of multiple Bmp4 expression sites upon
transition from refractory to competent telogen (L-N). Regain of Bmp4
expression in dermal papilla and secondary hair germ during anagen initiation
(O-Q). Blue color X-gal staining at the sites of Bmp4-lacZ transgene activity.
R-W) Temporal overlap of Bmp4 and Bmp2 in situ expression dynamics during
hair cycle.
Scale bars: A, H: 1 mm; B-E: 500 mkm; I-K, R-W, G, G: 200 mkm; F, F: 100
mkm; L-Q: 50 mkm.

226
Figure 4.3: Continued

227
Figure 4.3: Continued

228
Figure 4.3: Continued

229
We have further investigated the role of Bmp signaling during telogen
competence by studying expression patterns of the 52bpMsx2-hsplacZ transgene,
which acts as a faithful Bmp-responsive element (Brugger et. al, 2004). During
refractory telogen 52bpMsx2-hsplacZ is abundantly expressed in telogen hair
follicles (Figure 4.4D, 4.4F, 4.4J). Re-commencement of competent telogen is
associated with the downregulation of intra-follicular 52bpMsx2-hsplacZ
expression (i.e. loss of Bmp signaling activity; Figure 4.4E, arrowheads on 4.4G,
4.4K).

4.3.5 Resumption of Bmp signaling in anagen correlates with the loss of
propagation abilities by early anagen hair follicles
Bmp4 (Figure 4.3O-Q, 4.3T), Bmp2 (Figure 4.3B, 4.3W) and 52bpMsx2-hsplacZ
(Figure 4.4H, 4.4I) expression resumes in anagen and is maintained throughout
catagen into subsequent refractory telogen, closing the dynamic, intra-cutaneous
Bmp signaling loop. Interestingly, within the spreading wave, 52bpMsx2-hsplacZ
expression appears first in anagen V hair follicles and is absent in front of the
wave: from the telogen-anagen I wave front to anagen IV hair follicles (red
arrowheads on Figure 4.4I). This lack of Bmp signaling activity on the front end
of the spreading wave parallels the re-appearing pattern of Bmps. For example,
Bmp2 starts to be expressed first surrounding anagen III hair follicles of the
spreading wave (Figure 4.3B). It is not seen earlier (Figure 4.3B). The Bmp
antagonist noggin is expressed in the epithelial bulge of the hair follicles
throughout the hair cycle (based on NOG-lacZ transgene activity; Figure 4.4B).

230
Figure 4.4 Spatial and temporal changes of noggin expression and Bmp pathway
activity in hair follicles within a hair cycle domain.
A, B) Differential noggin expression throughout hair cycle stages as
demonstrated by NOG-lacZ transgene activity. Noggin is expressed in the hair
bulge area throughout the cycle. Mesenchymal noggin is expressed only during
anagen: first in the dermal sheath and later in the dermal papilla and basal stalk.
C) Schematic representation of expression sites and temporal changes of Noggin,
Bmp4 and Bmp2 expression relative to the morphological components of the hair
cycle domain and corresponding functional phases of the hair cycle.
D-I) Identification of Bmp pathway signaling within hair follicles during the hair
cycle. Bmp pathway signaling was based on differential 52bpMsx2-hsplacZ
transgene activity. Blue color (D-H) and green pseudocolor (I) X-gal staining at
the sites of Bmp4-lacZ transgene activity. Black arrow (G) and red arrow (I)
positive transgene expression in anagen hair follicles. Red arrowhead on G
absence of transgene expression from competent telogen hair follicles.
J, K) Localization of 52bpMsx2-hsplacZ transgene activity (blue colored X-gal
staining) within the companion cells compartment of club hairs derived from
refractory telogen hair follicles (J). Disappearance of transgene expression from
companion cells of competent telogen hair follicles (K). Black arrows point at
positive transgene expression sites.
Size bars: D, E: 500 mkm; F, G, I: 200 mkm; J, K: 20 mkm;
Bulge hair bulge; DP dermal papilla, DS dermal sheath; Stalk basal stalk.

231
Figure 4.4: Continued

232
Figure 4.4: Continued

233
Figure 4.4: Continued

234
However, expression of the mesenchymal NOG-lacZ is hair cycle-dependent.
NOG-lacZ is absent from the dermal compartment of all telogen hair follicles and
is expressed in the dermal papilla and basal stalk of anagen VI hair follicles. We
have identified the dynamical re-appearance of mesenchymal NOG-lacZ
expression within the anagen spreading wave. NOG-lacZ appears first throughout
the dermal sheath of anagen II hair follicles (Figure 4.4B). As hair follicles
progressively elongate during anagen IIIb-IV, expression of NOG-lacZ becomes
restricted to the proximal segment of dermal sheath and to the basal stalk. By
anagen VI, the proximal dermal sheath expresses NOG-lacZ very faintly. Most of
the NOG-lacZ expression is within the dermal papilla and basal stalk (Figure
4.4B). Thus, propagating anagen has the following Bmp signaling profile:
complete absence of Bmp2 expression, reappearance of some of Bmp4-lacZ
expression sites, strong dermal sheath expression of NOG-lacZ and lack of
52bpMsx2-hsplacZ transgene activity (Figure 4.1H, 4.4C). The Bmp signaling
profile of active anagen is as follows: strong Bmp2 expression, strong Bmp4-lacZ
expression in multiple sites, confinement of mesenchymal NOG-lacZ to the
dermal papilla and basal stalk at the very bottom of anagen hair follicles, and
strong 52bpMsx2-hsplacZ transgene activity (Figure 4.1H, 4.4C).

4.3.6 Skin-wide down-regulation of Bmp signaling in KRT14-NOG mice
shortens refractory telogen length and disrupts normal hair cycle domain
patterning
Our expression data suggests that refractory telogen is associated with high
intrafollicular Bmp signaling and that its downregulation results in the

235
recommencement of competence. To validate these findings we have
overexpressed ectopic noggin under the keratin 14 promoter in the KRT14-NOG
mouse. Within telogen, skin keratin 14 expression domains include the
interfollicular epidermis and almost the entire epithelial component of the hair
follicles. It surrounds the expression domain of the 52bpMsx2-hsplacZ transgene
(Figure 4.6A) and should oppose normal Bmp4 and Bmp2 signaling in telogen
hair follicles (Figure 4.6B).

Indeed, over-expression of noggin led to the effective down-regulation of Bmp
signaling activity in early telogen hair follicles, as judged by the premature loss
of 52bpMsx2-hsplacZ transgene expression in early telogen skin of KRT14-NOG
/ 52 bp 6Msx2-hsplacZ transgenic mice (Figure 4.6C). Furthermore, KRT14-NOG
mice have significantly shorter duration of refractory telogen and telogen
variability (Figure 4.5C). The length of telogen in dorsal KRT14-NOG hair
follicles is dramatically reduced (7.9 +/- 11 days (n=71)) with a minimal length
of only 6 days (vs. 28 days in WT) and a maximal length of only 11 days (vs. 88
days in WT). At the same time anagen length remains essentially unchanged
(12.3 +/- 1 days (n=71) vs. 12.7 +/- 0.97 days (n=18) in WT). It appears that in
KRT14-NOG mice, competence to respond to growth stimulating signals is
restored only 6 days after the previous anagen/catagen ends. Similar changes in
telogen timing dynamics are found in the ventral skin of KRT14-NOG mice.
Here, the length of anagen remains largely unchanged (7.1 +/- 0.8 days (n=55)
vs. 7.6 +/- 0.7 days (n=28) in WT). However the duration of telogen is
dramatically reduced (8.9 +/- 1 days (n=41)) with minimal length of only 6 days

236
(vs. 25 days in WT) and maximal length of only 11 days (vs. 78 days in WT). As
a result of accelerated regain of telogen competence, KRT14-NOG mice show an
increased rate of hair growth cycling turnaround. KRT14-NOG hair follicles also
display an altered response to hair plucking. The differences in plucking-induced
anagen timing seen in WT mice are eliminated in KRT14-NOG mice (Figure
4.5D, 4.5E). In all cases skin areas with either 50 or 200 club hairs plucked on
day 1 of new telogen re-entered new anagen 6-7 days afterwards (vs. at least 28
days in WT) together with surrounding telogen skin where hair follicles were not
stimulated by plucking.

Over-expression of ectopic noggin in KRT14-NOG mice also should prevent the
early loss of propagation abilities by anagen hair follicles and should disturb
border stabilization. Indeed, KRT14-NOG mice do not display stable domain
borders, but rather show continuous movement of the wave-front of hair growth
domains. Largely simplified patterns of hair growth domains arise (Figure 4.5A,
4.5B).

4.3.7 Bmp signaling activity and telogen timing in KRT14-NOG hair follicles
can be partially restored by a normal skin microenvironment
To test whether hair cycle differences in KRT14-NOG mice are caused by
reduced Bmp signaling during telogen or by permanent developmental changes to
the hair follicles we performed transplantation of a small group of KRT14-NOG
hair follicles (~100 hair follicles) into the adult SCID mouse skin. This changes
the typical growth behavior of KRT14-NOG hair follicles (Figure 4.6D).

237
Figure 4.5 Overexpression of noggin leads to early restoration of telogen
competence and disruption of hair growth patterns.
A, B) Simplification of hair cycle domain patterns in KRT14-NOG mice.
KRT14-NOG mice do not display stable domain borders, but rather show
continuous movement of the wave-front of hair growth domains.
C) Timing of telogen length in KRT14-NOG mice reveals a shortening of the
refractory phase. D) Differences in plucking-induced anagen timing are
eliminated in KRT14-NOG mice as a result of accelerated regain of telogen
competence. Red arrowheads point at hair growth sites within the regions of
plucking. E) Comparison of response time of refractory telogen skin to club hair
plucking stimulation between WT and KRT14-NOG mice.

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Figure 4.6 Interactions of KRT14-NOG skin transplant with the host SCID skin.
A, B) Overlapping domains of 52bpMsx2-hsplacZ transgene activity (blue) and
keratin 14 expression (brown; A). Schematic illustration of Bmp2 (purple), Bmp4
(red) and transgenic noggin (green) expression domains within refracory telogen
skin of KRT14-NOG mouse (B).
C) Premature loss of 52bpMsx2-hsplacZ transgene activity in early telogen hair
follicles of KRT14-NOG / 52 bp 6Msx2-hsplacZ transgenic mice. The blank
arrow points to the direction of the telogen propagation (i.e. anagen regression).
D) Partial restoration of growth modes within small groups of KRT14-NOG hair
follicles (~100 hair follicles) transplanted into the SCID skin microenvironment.
Blank arrow advancement of the spreading wave; yellow dotted line
wavefront.
E) Preservation of fast anagen re-entry within large groups of KRT14-NOG hair
follicles and induction of early competence in SCID hair follicles along the ~ 3
mm perimeter of KRT14-NOG skin transplant. Green dotted line border
between anagen and telogen hair follicles of SCID host.
F, G) Restoration of 52bpMsx2-hsplacZ transgene activity during early telogen
within small groups of KRT14-NOG / 52 bp 6Msx2-hsplacZ hair follicles (~100
hair follicles) transplanted into the SCID skin microenvironment. Blue color
X-gal staining at the sites of 52bpMsx2-hsplacZ transgene activity. Yellow arrow
- transgene expression in early telogen hair follicles.
H) Restoration of telogen timing within small groups of KRT14-NOG hair
follicles (~100 hair follicles) transplanted into the SCID skin microenvironment.
Scale bars: C: 500 mkm; A, F, G: 50 mkm

239
Figure 4.6: Continued

240
Figure 4.6: Continued

241
Figure 4.6: Continued

242
Instead of accelerated regain of telogen competence and swift re-initiation of
anagen following short telogen, transgenic hair follicles partially synchronize
their cycles with the surrounding host SCID skin. They can remain in telogen for
an extended period of time and respond to an anagen spreading wave originating
from the host's initiation center. At the same time, groups of KRT14-NOG hair
follicles retain a higher propensity for spontaneous anagen initiation, resulting in
growth which is out-of-phase with still refractory host skin or in spontaneous
growth within the groups, followed by the induction of the surrounding
competent host skin. This indicates partial rescue of the hair cycling behavior.

As part of the small group surrounded by the host SCID skin, KRT14-NOG hair
follicles spend more time in telogen (an average of 38 days vs. 7.9 days as part of
the donor KRT14-NOG skin; Figure 4.6H). Additionally, the range of telogen
timing for KRT14-NOG hair follicles expands from very narrow (6 to 11 days) to
significantly wider (11 to greater than 100 days). Telogen timing within a small
group of transplanted KRT14-NOG hair follicles is compatible to the telogen
timing observed for the small group of transplanted control (C57BL/6J) hair
follicles (averaging 42 days with the distribution from 13 to greater than 100
days).

Transplantation of a small group of KRT14-NOG / 52 bp 6Msx2-hsplacZ hair
follicles into adult SCID mouse skin prevents the early loss of Bmp signaling
activity within transgenic hair follicles as judged by the retention of
52bpMsx2-hsplacZ transgene expression in early telogen hair follicles (Figure

243
4.6G: Tel). Anagen and catagen KRT14-NOG / 52 bp 6Msx2-hsplacZ hair
follicles show typically strong 52bpMsx2-hsplacZ transgene expression in
various compartments of the hair follicle (Figure 4.6G: An, Cat vs. 4.6F: An,
Cat). Transplantation of a small group of control, 52 bp 6Msx2-hsplacZ hair
follicles (~100 hair follicles) into adult SCID mouse skin does not alter typical
52bpMsx2-hsplacZ transgene expression during early telogen (Figure 4.6F: Tel)
seen in donor 52 bp 6Msx2-hsplacZ transgenic mouse skin.

Next, we transplanted a large group of KRT14-NOG hair follicles (skin circle
1cm in diameter) into adult SCID mouse skin to see if this can alter the typical
growth behavior of SCID hair follicles on the borderline with KRT14-NOG skin
(Figure 4.6E). Indeed, transplanted KRT14-NOG hair follicles continue to
re-enter anagen after only a short period of telogen. At the same time, telogen
SCID hair follicles along the perimeter of KRT14-NOG skin (2.8 mm) acquire
early competence (only on the 11th day of telogen) and re-enter new anagen
together with KRT14-NOG hair follicles. These hair follicles gain competence
despite the fact that the rest of the SCID hair follicles further away from the
KRT14-NOG transplant remain in refractory telogen.

Transplantation data suggests that telogen competence of both KRT14-NOG and
SCID hair follicles can be modulated through changes in the BMP signaling
balance. The increase of refractory telogen length in small groups of
KRT14-NOG hair follicles was achieved by diluting the strength of the
transgenic noggin upon transplantation into SCID skin. The shortening of

244
refractory telogen in SCID hair follicles was achieved due to high amounts of
transgenic noggin leaking out of the edge of the large group of transplanted
KRT14-NOG skin.

4.3.8 Recapitulation of hair cycle patterns with cellular automata model
In order to recapitulate the hair cycle patterning behavior observed in mouse skin
we have used a cellular automata model (REF). We have chosen this model
because it satisfies several important requirements: a) it allows us to model each
individual hair follicle as a single automaton and multiple hair follicles as a
two-dimensional lattice of multiple automata; b) it us allows to model hair cycle
stages as a set of consecutive states that a single automaton cycles through; 3) it
allows us to model the interactions between hair follicles and the interfollicular
microenvironment as interactions between multiple automata which are
determined by a combination of stochastic and deterministic factors. We have
introduced 16 different parameters to describe the cellular automata model
behavior. Several parameters and their values were chosen to simulate the
strength and duration of Bmp signaling during different stages of the hair cycle.
See Supplement 2 for detailed information on the logic behind the model and
technical aspects of the modeling process. Upon graphical simulation of the
model a number of interesting phenomena are observed which correlate to those
observed on the mouse skin: wave spreading, formation of initiation centers and
boundaries. We have simulated spreading of a wavefront of hair follicles in
propagating anagen (P) over a competent telogen (C)

245
Figure 4.7 Functional phases of the hair cycle and recapitulation of hair cycle
patterns with cellular automata model.
A) Schematic representation of new functional stages of the hair cycle (refractory
telogen, competent telogen, propagating anagen and active anagen) overimposed
on morphological stages of the hair cycle (telogen, anagen and catagen) and
expression dynamics of dermal noggin, and Bmp signaling activity. B)
Simulation of spreading of a front of propagating anagen (P) hair follicles over a
competent telogen (C) field. C) Simulation of formation of multiple initiation
centers on the competent telogen field. D) Simulation of visible boundaries
between different follicle states.

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Figure 4.7: Continued

247
Figure 4.7: Continued

248
Figure 4.7: Continued

249
field (Figure 4.7B). Initially, all follicles are set to be in C phase, with a certain
fraction having the ability to spontaneously enter P phase. Once a critical number
of hair follicles has spontaneously entered P phase together, they form a new
initiation center (grey spot on Figure 4.7B at 14.8 days). As time increases,
further follicles in C phase are able to enter P phase due to the presence of
supra-threshold stimuli from follicles already in P phase (grey area on Figure
4.7B at 19.8 days). Sustained stimuli from cells in P ensure that the wave
continues to propagate over the competent follicle field, until all follicles have
entered P phase. With the course of time, hair follicles across the entire patterning
field have cycled through P → A → R and start to become competent again
(white area on Figure 4.7B at 41.4 days). Upon gain of competence, the initiation
of a new wave begins again once enough follicles have become able to
spontaneously enter P phase and form a new initiation center (black spot with
grey halo on Figure 4.7B at 74.7 days).

We have also simulated formation of multiple initiation centers by slightly
increasing the probability of spontaneous entry from C to P phase (Figure 4.7C).
As multiple initiation centers form (Figure 4.7C at 11.5 days), wave-fronts
originating from these centers eventually collide and merge (Figure 4.7C at 14.8
days). After an initial round of cycling, the patterning field becomes competent
once again and a further set of initiation centers forms (Figure 4.7C at 71.4 days).
The arising simulation patterns resemble natural patterns formed by several
initiation centers.

250
Boundary formation between hair cycle domains can be simulated when different
areas of the patterning field begin to oscillate out of synchrony (Figure 4.7D;
here, the bottom half of the field is just entering the R phase, whilst the upper half
is in C phase). Although initiation centers form in the upper half of the field, the
resulting wave is unable to propagate to lower regions which are still in R phase
(Figure 4.7D at 18.1 days). On the other hand, once the lower half of the field
enters C phase (Figure 4.7D at 24.8 days), there is not a sufficient number of hair
follicles in P phase remaining in the upper half to stimulate entry to P phase.
Thus follicles in the lower half of the field must rely on the formation of new
initiation centers (Figure 4.7D at 36.4 days). This process repeats with the upper
field on the subsequent cycle (Figure 4.7D at 73.0 days). However, it is possible
that the entire field will re-synchronize if there are follicles in the upper half of
the field still able to stimulate entry to P phase by the time follicles in the lower
enter C phase. This simulation closely resembles multiple accounts of border
stabilization as well as the merging of domains observed in mice.

4.4 Discussion
4.4.1 We have initiated this study because we were intrigued by the patterned hair
growth observed in Msx2
-/-
mice with cyclic alopecia (Ma et al., 2003), in nude
mice with transversing waves phenotype (Suzuki et al., 2003) as well as in
normal mice and rats described in the classical literature on skin (Durward and
Rundall, 1949; Mottram, 1945). We found that patterns result from coordinated
transition between hair cycle stages among hair follicle populations.
Telogen-anagen transitions or lack thereof are critical to the pattern formation

251
process. They determine the size, shape and physical boundaries between hair
cycle domains in two alternative stages of hair cycle activity: growth (i.e. anagen)
and quiescence (i.e. telogen).

Our results indicate that the period of quiescence (i.e. telogen itself) preceding
transition into anagen consists of two new phases: refractory and competent.
Telogen begins with a refractory phase (i.e. minimal telogen). It is characterized
by the inability of hair follicles to re-enter anagen. This is manifested both by the
inability to enter anagen spontaneously (i.e. form initiation centers) or in response
to anagen spreading waves from neighboring skin. Pioneer studies of patterned
hair growth have implied the existence of this phenomenon, although this line of
investigation has not been pursued (see Introduction; Durward and Rundall,
1949; Ebling and Johnson, 1961). Later it was suggested that so-called chalone(s)
- substances inhibiting anagen development - are present in telogen skin (Paus et.
al, 1990). Our work uncovers the molecular basis of the phenomenon of
refractory telogen and places it at the center of the local paracrine mechanism of
hair cycle control. Our results indicate that in WT (C57BL/6J) mice refractory
telogen persists for about 4 weeks, whereupon follicles enter a competent phase.
During this phase they can either enter anagen spontaneously or respond to an
anagen spreading wave. On some occasions hair follicles re-enter anagen
immediately after refractory telogen ends. However, in many cases they remain
in competent telogen for a variable length of time (anywhere from 0 to 60 days in
our experiments). The exact duration of competent telogen is subject to variation
and depends on the availability of anagen spreading signals in the vicinity of the

252
follicle or on the stochastic nature of formation of initiation center(s). If anagen
spreading occurs in a region adjoining the domain that has just acquired
competence it will spread into this domain.

However, not all anagen hair follicles can propagate anagen initiation signals to
competent telogen hair follicles with the same level of efficacy. While early
anagen hair follicles at the tip of the spreading wave are very good at it, late
(anagen VI) hair follicles are not. On this basis we distinguish between two
functional phases of anagen: propagating (early) and active (late anagen VI). The
inefficacy of propagation displayed by anagen VI hair follicles constitutes the
phenomenon of boundaries stabilization, whereas boundaries between active
anagen and competent telogen do not shift. We conclude that hair cycle domain
patterns depend on the relative distribution of hair follicles in all four functional
stages of the hair cycle on the surface of the skin and the time it takes to restore
telogen competence and lose propagation ability in the anagen phase.

Our work uncovers a new role for Bmp signaling in the maintenance of refractory
telogen and termination of propagating anagen. Complex new extra- and
intrafollicular expression dynamics of Bmp2/4 and noggin appear to regulate
these events. Our experiments confirm that skin-wide down-regulation of Bmp2/4
expression is responsible for the gain of telogen competence. Refractory telogen
is characterized by multiple extra- and intrafollicular sites of Bmp2 and Bmp4
expression. Together they create a continuous high-Bmp signaling
microenvironment surrounding telogen hair follicles. Domains of high Bmp

253
signaling can be clearly visualized by whole mount staining techniques. They
form distinguishable boundaries with domains in competent telogen that are
mostly void of Bmp2 and Bmp4 expression sites. Our results suggest that Bmp2
and Bmp4 act synergistically during the refractory phase and disappear around
the same time during the regain of competence. Largely decreased Bmp signaling
activity on the borderline of competent telogen and propagating anagen is an
essential prerequisite of successful anagen spreading. Our results show that
within the spreading wave both Bmp2 and Bmp4 expression and Bmp signaling
activity do not resume until hair follicles reach advanced stages of anagen.
Moreover, whilst low in Bmp2 and Bmp4, the leading segment of the anagen
spreading wave (i.e. propagating anagen) has a high content of the Bmp
antagonist, noggin, expressed throughout the dermal sheath of early anagen hair
follicles. Detailed whole mount analysis of NOG-lacZ expression during all hair
cycle stages shows constitutive rather than on-off expression of noggin in the
epithelial bulge (Zhang et. al, 2006) and suggests that the dermal sheath is the
primary source of noggin driving the level of Bmp signaling even further down
during early anagen. Thus, the actual telogen-to-anagen transition and the
progression through early stages of anagen occur under the condition of virtual
absence of Bmp signaling activity. As anagen progresses, all Bmp2 and Bmp4
expression sites rapidly re-appear and mesenchymal noggin expression becomes
confined to the dermal papilla and basal stalk at the very bottom of elongated
anagen hair follicles, away from the skin surface. Regain of Bmp signaling
activity ends the propagation phase of anagen and contributes to the phenomenon
of boundary stabilization.

254
Previously, Bmp signaling was shown to control hair follicle development (Lyons
et. al, 1989; Noramly and Morgan, 1998; Botchkarev et. al, 1999; Botchkarev et.
al, 2002; Botchkarev, 2003; Plikus et. al, 2004), differentiation of progenitor cells
and hair keratin expression in postnatal hair follicles (Kulessa et. al, 2000;
Kobielak et. al, 2003; Yuhki et. al, 2004; Andl et. al, 2004), as well as catagen
regression (Botchkarev, 2003; Andl et. al, 2004; Guha et al., 2004; also see
review by Botchkarev and Sharov, 2004). Additionally, Bmp signaling was
shown to be crucial for new anagen initiation (Botchkarev et al., 2001). Our
new data allows us to extend the existing knowledge of the involvement of Bmp
signaling in the hair cycle and to explain the phenomenon of patterned
distribution of hair cycle stages. According to our model, Bmp signaling controls
two critical points of the hair cycle. First is the preceding event of
refractory-to-competent telogen transition. Second is the point of
telogen-to-anagen transition. Previous research has conclusively shown that
telogen-to-anagen transitions can be promoted by adding extraneous Bmp
antagonist, noggin, into the skin. Following the anagen initiation event,
endogenous mesenchymal noggin starts to be expressed in early anagen hair
follicles (Botchkarev et al., 2001). This is accompanied by down-regulation of
Bmpr-IA in the secondary hair germ which ensures that newly initiated anagen
will proceed, rather than being terminated by a sudden rise in Bmps. Indeed, once
initiated under favorable physiological conditions, anagen continues without
interruption until its completion.

255
However, recognition only of the telogen-to-anagen transition does not allow one
to explain why some telogen hair follicles (early telogen) expressing Bmpr-IA
respond to anagen spreading signals (such as noggin) and others (late telogen hair
follicles) do not. It does not explain why and how this ability is regained and also
why this it occurs after a certain period of time and simultaneously over large
groups of hair follicles (i.e. entire hair cycle domains). Here we have shown that
timed downregulation of Bmp2 and Bmp4 expression during telogen precedes the
event of new anagen induction and constitutes the event of regain of telogen
competence. Low Bmp signaling activity in competent telogen hair follicles
creates a favorable microenvironment for successful anagen initiation, when
noggin, as has been previously suggested and also shown in this work, can play
the role of an anagen promoting agent by further reducing Bmp activity below the
threshold level required to keep secondary hair germ cells from proliferating (i.e.
engaging into an active anagen; Botchkarev et al., 2001). The introduction of a
refractory-to-competent telogen transition point provides explanation as to why,
even in the presence of an anagen spreading wave (i.e. source of secreted anagen
inducers, such as noggin), hair follicles from the refractory telogen domain
remain unresponsive. The amount of Bmp2 and Bmp4 produced within the
refractory domain simply overpowers small amounts of noggin generated within
the spreading wave.

That the length of refractory telogen is subject to dynamic regulation by the Bmp
signaling pathway rather than being a genetically predetermined property of hair
follicles is supported by several lines of evidence. Advancement of competence

256
can be experimentally facilitated by supplying large amounts of extraneous Bmp
antagonist. Overexpression of noggin in the KRT14-NOG mice used in this study
or in NSE-Noggin mice (Guha et al., 2004), as well as the delivery of extraneous
noggin upon intracutaneous implantation of noggin-soaked beads, (Botchkarev et
al., 2001) drives Bmp signaling activity down, advancing competent telogen.

Further, saturation of telogen skin with the supplementary noggin simulates the
role of physiological noggin and causes premature anagen induction. The
duration of refractory telogen can be either experimentally lengthened for
KRT14-NOG hair follicles or shortened for WT hair follicles upon transplantation
of groups of KRT14-NOG hair follicles of variable size into WT (SCID) skin. If a
small group of KRT14-NOG hair follicles (~ 100 hair follicles) is transplanted,
the effect of noggin is neutralized by excessive amounts of Bmps entering into
the transplant from the surrounding area. Consequently, the length of refractory
telogen within the transplant increases. If a large piece of KRT14-NOG skin is
transplanted (~ 1x1 cm), normal levels of Bmp signaling on the host side of the
donor-host skin boundary are reduced by transgenic noggin leaking out of the
transplant. The length of refractory telogen for the WT host hair follicles along
the perimeter of the KRT14-NOG transplant decreases. Transplantation of WT
hair follicles does not alter the duration of refractory telogen. Dynamic changes
in telogen competence can also be revealed upon implantation of Bmp4-soaked
beads into WT telogen skin. Saturation of telogen skin with extraneous Bmp4 can
simulate reversal of competent telogen into refractory telogen and can prevent
development of hair plucking-induced anagen (Botchkarev et al., 2001). This also

257
agrees with our results where significant delays in hair-plucking induced anagen
were observed in refractory over competent telogen skin. Localized experimental
manipulations of the Bmp balance (via bead implantation and KRT14-NOG hair
transplantation) can cause local disturbances in hair cycle domain patterns of the
host. Overexpression of noggin throughout the entire skin (in KRT14-NOG mice
themselves) causes global changes in the normal patterning process. Due to the
accelerated regain of telogen competence in KRT14-NOG mice,
anagen-refractory telogen boundaries virtually never form. Stable hair cycle
domains, as we observe them in WT mice, are absent. Instead, KRT14-NOG mice
show successive waves of anagen sweeping across the skin.

In order to test if our assumptions regarding the hair cycle patterning mechanism
are correct and if dynamic changes in Bmp signaling over the course of the hair
cycle are sufficient to support this patterning behavior we have simulated our
experimental data using a cellular automata model. In order to conform our data
to the logic of the model each hair follicle is described by one automata whose
behavior is determined by a combination of stochastic and deterministic factors.
Stochastic effects come into play when determining which competent telogen (C
phase) hair follicles become spontaneously able to enter propagating anagen (P
phase). On the other hand, hair follicle behavior has a deterministic component:
every follicle must cycle in order through successive states (hair cycle stages),
and if competent, a follicle must re-enter P phase given a sufficient stimulus from
neighboring P phase follicles. Taken together, neighbourhood rules introduced
into the model simulate the effect of dynamic Bmp signaling on the functional

258
status of hair follicles and transition events in between phases: refractory telogen
→ competent telogen and competent telogen → propagating anagen. Because our
model is parameterized experimentally it is able to display the key patterning
phenomena observed on the skin: formation of initiation centers, propagation of
the anagen wave and formation of the boundary. The modeling results suggest
that our assumptions on the patterning mechanism are correct in principal, and
that dynamic changes in Bmp signaling and the effect it exerts on the functional
status of hair follicles constitute the minimal set of requirements needed for the
patterning process to perpetuate.

In our work we have identified a conceptually new mode of hair cycle control.
Rather than being an autonomous intra-follicular event, the telogen-anagen
transition is largely regulated by signals originating from the extra-follicular
environment and neighboring hair follicles. We have identified the important role
that various extra- and intra-follicular, largely mesenchymal Bmp2, Bmp4 and
noggin, sources play on intra-follicular Bmp signaling output during telogen,
immediately prior to, and during, the event of telogen-anagen transition. Newly
gained knowledge allows us to view the hair cycle in the content of the
microenvironment surrounding the hair follicle. It allows us to explain how hair
follicles make decisions on hair cycle progression by listening to other follicles
and inter-follicular signaling centers and how multiple hair follicles can
coordinate their hair cycles to form macroscopic growth patterns. Our work
opens new area for hair cycle control studies that takes an integrative approach
and views hair follicles in relation to the surrounding stroma and neighboring

259
follicles. New studies should recognize the existence of refractory and competent
phases of telogen and the ability of early anagen hair follicles to propagate
activation signals to competent telogen follicles. Studying the dynamics of other
signaling pathways throughout telogen phases and immediately prior to anagen
induction should help to elucidate the molecular events that precede and cause
initiation of anagen rather than regulate early post-induction events.

260
Conclusion
In this work I investigate complex morphoregulatory role of Bmp signaling
pathway in ectodermal organs development and post-developmental cyclic
growth activity. Concept of morphoregulation suggests that mild spatial-temporal
changes in signaling pathway(s) activity during organs development can
modulate its morphological phenotype. Morphoregulation provides a plausible
mechanism of how structure of organs can adjust to fulfill their function or
assume new function under changing conditions, and how phenotypical
diversification of organs can arise between different species. Yet, experimental
testing of morphoregulatory functions of major signaling pathways is
challenging. Many available transgenic mouse systems display severe
pathological defects or even failure of organs development. In order to suggest a
possible morphoregulatory function for the signaling pathway one must study
phenotypical changes that fall into the category of physiological or
semi-physiological.

In my work I have concentrated on studying phenotypical changes of ectodermal
organs because of the multiple advantages that they have over other experimental
systems. First of all, in laboratory settings many functions of ectodermal organs
are dispensable for animals survival. Thus, our experimental system can allow
presence of both physiological as well as pathological phenotypes without
causing lethality. Secondly, there are many types of ectodermal organs on the
surface of the animal and many of them exist in multiple copies (such as teeth,
hair follicles, claws etc). Phenotypes of multiple organs can be more insightful

261
over the phenotype of one organ only. And lastly, phenotipical changes of
ectodermal organs are easy to identify and observe due to their accessibility on
the surface of the animal. In order to generate mostly physiological,
semi-physiological or mild pathological phenotypes of ectodermal organs we
have chosen to use a transgenic mouse system where Bmp antagonist is
dynamically expressed under specific ectodermal promoter, rather than entirely
abolishing Bmp signaling via loss-of-function mutations. We have succeeded in
our goal by producing K14-noggin transgenic mouse, where Bmp antagonist
noggin is expressed under keratin 14 promoter specific to epidermis, oral mucosa
epithelium, as well as to the derivative ectodermal structures. By studying both
low and high K14-noggin transgene copy number mice we have obtained a
wealth of phenotypes detailed in appropriate sections of this thesis. Taken
together phonotypical variations of ectodermal organs of K14-noggin mice
demonstrate morphoregulatory role of Bmp signaling pathway during all stages
of development: induction, morphogenesis and differentiation. However, one
limitation of our system is in inability to attribute a particular morphoregulatory
function of Bmp signaling to some of the similar phonotypical variations of
ectodermal organs seen among different animals. For example, our studies
suggest that Bmp signaling controls pattering of the molars crown and roots. It is
yet to be established if spatial-temporal variations of Bmp signaling produce
similar molars phonotypes in natural settings -in mammals other than
laboratory mice. New experimental models and methods are required to study
signaling pathways behind phenotypical diversity of ectodermal organs among
various animals. Additionally, exact mechanisms of spatial-temporal changes of

262
signaling pathways activities in natural settings can differ from these induced by
us and others in transgenic mouse systems. Control can be exerted on the level of
promoters, transcription, splicing, translation, and multiple post-translational
modifications of various signaling pathways agonists, antagonists, receptors and
downstream targets.

In order to address this apparent limitation of our experimental system I have
investigated physiological mechanism behind the phenomenon of patterned hair
growth. Upon completion of their development hair follicles engage into the
cyclic regenerative activity, known as the hair cycle. Because of the similarity of
the hair cycle to the hair follicle morphogenesis, hair cycle can be viewed as the
direct extension of morphogenesis into adulthood. While majority of previous
studies aimed at investigating intra-follicular control of hair cycle, we have
investigated the role of extra-follicular environment on hair cycle within
individual hair follicle. For the first time I have conclusively showed that
extra-follicular environment plays major role in physiological regulation of hair
cycle in large populations of hair follicle. Through the series of experiments
detailed in Chapter 4 of this thesis I have conclusively demonstrated
physiological role of Bmp signaling pathway in the process of inter-follicular hair
cycle regulation. Furthermore I have identified detailed mechanism of Bmp
signaling activity regulation throughout the hair cycle. I have shown that Bmp
signaling activity is modulated largely by spatial-temporal changes of expression
levels of Bmp agonists - Bmp2 and Bmp4, and Bmp antagonist noggin in
multiple extra- and intra-follicular tissue compartments within the skin. I hope

263
that this study will become a cornerstone work in the series of follow-through
studies detailing the role of extra-follicular environment and involvement of other
signaling pathway in physiological regulation of hair cycle among large
populations of hair follicles. As the direct continuation of this study I plan to try
to identify a potential clock mechanism controlling timed expression of Bmps and
noggin during hair cycle. In addition I am undertaking an ongoing effort to
extend our findings beyond the mouse system. I have already demonstrated that
rabbits exhibit similar patterning mode of hair cycle as mice do. However,
multiple differences exist between hair growth patterns in rabbits and mice. In
general, rabbits form patterns of much higher complexity. I am working on
identifying principal differences in hair cycle patterning process between these
two mammalian species. Further research efforts will be directed toward
identification of differences in Bmp signaling among these to species and how
they can translate into hair cycle pattern differences. At the same time, some
mammalian species, such as guinea pigs and humans lack patterned hair cycle
mechanism. In guinea pigs every hair follicle appears to cycle independent of its
neighbors and hair cycle patterns never form. Preliminary transplantation of
guinea pigs skin into mouse host failed to restore patterned hair growth among
guinea pigs hair follicle. This suggests that patterned hair growth mechanism
regulated by Bmp signaling pathway is turned-off intrafollicularly. My future
studies will be directed toward identification of the shut off point that allows
guinea pigs hair follicles to escape patterned hair growth regulation mechanism.

264
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Abstract (if available)

Abstract Ectodermal organs are produced through the series of epithelial-mesenchymal interactions followed by topological transformation of the otherwise flat ectodermal layer. Fundamental signaling pathways regulate development of ectodermal organs. Fine tuning of the organ's size and shape during development is achieved by mild spatial-temporal changes of signaling pathway activity rather than on-off switches. In my work I have investigated the extent of morphoregulatory activities of Bmp signaling pathway. I have modulated normal Bmp signaling by overexpressing of Bmp antagonist noggin under keratin 14 promoter in K14-noggin mice. I have shown that changes in Bmp pathway activity can alter various ectodermal organs at different developmental stages: induction (increased number of pelage hair follicles, formation of compound vibrissa follicles, claw agenesis, transdifferentiation of sweat glands into hair follicles etc.), morphogenesis (defect of eyelids opening, enlargement of external genitalia etc.) or differentiation (incomplete differentiation of claw plates, retarded differentiation of hairy spines). I have further shown that in morphologically complex teeth, Bmp pathway regulates all of the above stages of development.

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University of Southern California Dissertations and Theses

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Asset Metadata

Creator Plikus, Maksim V. (author)

Core Title Morphoregulation and cycling of ectodermal organs

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Pathobiology

Publication Date 04/19/2007

Defense Date 11/30/2006

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

Tag BMP,hair follicle,Noggin,OAI-PMH Harvest,stem cells

Language English

Advisor Chuong, Cheng-Ming (committee chair), Bellusci, Saverio (committee member), Jiang, Tingxin (committee member), Maxson, Robert E. (committee member), Tuan, Tai-Lan (committee member)

Creator Email plikus@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m423

Unique identifier UC1131383

Identifier etd-Plikus-20070419 (filename),usctheses-m40 (legacy collection record id),usctheses-c127-482153 (legacy record id),usctheses-m423 (legacy record id)

Legacy Identifier etd-Plikus-20070419.pdf

Dmrecord 482153

Document Type Dissertation

Rights Plikus, Maksim V.

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