University of Florida Thesis Or Dissertation Formatting

MOLECULAR INTERACTIONS BETWEEN THE ZPA AND THE AER IN THE DEVELOPING VERTEBRATE LIMB

CORTNEY MICHAEL BOULDIN

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2010

To my wife and family

ACKNOWLEDGMENTS

Without the moral and material support of my wife and family, this work would have been impossible. For all of their love, I am eternally grateful. I offer a special thank you to my wife, Erin DeFries Bouldin, both for allowing me the space to pursue dissertation work and for tutoring me in the subtleties of statistical analysis.

I thank my mentor Dr. Brian Harfe for introducing me to the fascinating field of developmental genetics. Dr. Harfe’s enthusiasm and unique ability to provide both support and freedom have been invaluable throughout my graduate career. I am also grateful to the members of my advisory committee of Dr. Martin Cohn, Dr. Jorg Bungert,

Dr. S. Paul Oh for their guidance and support.

Innumerable teachers throughout my education have inspired me to follow a career in science. In particular, I thank Mr. Terry Nusbaum for instilling an early interest in the biological sciences, and Drs. Brian Cain and Thomas Yang for introducing me to the wonders of laboratory science outside of a classroom.

I thank all the members of the Harfe Lab, including but not limited to Jason Rock,

Danielle Maatouk, Kyunsuk Choi, Jennifer Maier, Kendra McKee, Ben Cole, John Palsis and Bhavana Vangara, for friendship, technical assistance and countless helpful discussions. I thank the members of the Cohn lab for friendship and helpful discussion.

Outside of the University of Florida, I thank Dr. Amel Gritli-Linde, Dr. William Scott, Jr.,

Matthew McFarlane, Dr. Steve Vokes and Dr. Andrew McMahon for conversations and sharing unpublished data, which helped to shape the progress of this work. Finally, I am indebted to Joyce Connors, Michelle Ramsey and Jenneene Spencer for their outstanding ability to manage the practical aspects of graduate work and the staff at

Morningside Nature Center for access to their Dorking chickens.

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 7

LIST OF FIGURES ...... 8

ABSTRACT...... 10

CHAPTER

1 THE MOLECULAR BASIS OF LIMB PATTERNING...... 12

Sustained Outgrowth of the Vertebrate Limb ...... 13 The Apical Ectodermal Ridge and the Progress Zone Model ...... 13 A Molecular Model for Limb Outgrowth ...... 15 Patterning within a Developing Vertebrate Limb ...... 17 The Zone of Polarizing Activity (ZPA) and Molecules Expressed within the ZPA ...... 17 Mechanisms of Action for Shh ...... 17

2 NETWORKS OF SIGNALING WITHIN THE DEVELOPING LIMB ...... 21

Establishment of the AER ...... 21 Establishment of the ZPA ...... 21 Interactions between the ZPA and the AER ...... 22 Roles for Gene Expression Networks within the Developing Limb ...... 23

3 THE SHH SIGNALING PATHWAY IS PRESENT AND REQUIRED WITHIN THE VERTEBRATE LIMB BUD APICAL ECTODERMAL RIDGE FOR NORMAL AUTOPOD PATTERNING ...... 25

Results ...... 27 SHH Protein and Components of the Hedgehog Signaling Pathway are Present in the AER ...... 27 Hedgehog Signaling in the AER is Required for Formation of a Normal Autopod ...... 30 AER Length is Controlled by Hh Signaling within the AER ...... 31 Hedgehog Signaling in the AER Regulates the Shh/Grem1/Fgf Feedback Loop ...... 32 Discussion ...... 33 Experimental Procedures ...... 36 Mouse Alleles and Breeding ...... 36 Fate Mapping, β-Galactosidase Detection and Immunohistochemistry ...... 36 Whole Mount RNA in situ Hybridization and AER Measurement ...... 37

Bead Implantation and LysoTracker Analysis ...... 38

4 ABERRANT FGF SIGNALING, INDEPENDENT OF ECTOPIC HEDGEHOG SIGNALING, INITIATES PREAXIAL POLYDACTYLY IN DORKING CHICKENS .. 54

Introduction ...... 54 Results ...... 56 Dorking Chickens have Partially Penetrant Preaxial Polydactyly in the Hindlimb but not in the Forelimb...... 56 Ectopic Hedgehog Signaling Occurs in the Anterior of Late Stage Dorking Hindlimbs ...... 57 Gene Expression in the AER is Expanded in Dorking Hindlimbs ...... 58 Ectopic Fgf4 Precedes Ectopic Shh in Dorking Hindlimbs ...... 58 Extended Expression of Fgf4 in the Dorking Hindlimb is Maintained Independent of Hedgehog Signaling...... 59 Cell Death is Decreased in the Anterior Necrotic Zone of Dorking Hindlimbs ... 60 Inhibition of Ectopic Fgf but not Hedgehog Signaling in the Dorking Hindlimb can Rescue the Reduction of Cell Death Present in the Anterior Necrotic Zone ...... 61 Discussion ...... 62 Multiple Roles for FGFs in the Formation of Supernumerary Digits in Dorkings ...... 62 The Role of Shh in Dorking Polydactyly ...... 64 Potential Genetic Causes for Dorking Polydactyly ...... 66 Experimental Procedures ...... 66 Embryo Collection ...... 66 Skeletal Preparation ...... 67 Whole Mount RNA in situ Hybridization ...... 67 RNA Isolation, cDNA Synthesis and RT-PCR ...... 68 LysoTracker and Drug Treatments...... 68

5 CONCLUDING REMARKS ...... 78

APPENDIX

A CRE RECOMBINASE EXPRESSION DRIVEN BY THE PEAK7 ELEMENT OF THE PTCH1 PROMOTER ...... 83

B OLIGONUCLEOTIDES USED AS GENOTYPING PRIMERS ...... 88

C PROBES USED FOR RNA IN SITU HYBRIDIZATION ...... 90

LIST OF REFERENCES ...... 91

BIOGRAPHICAL SKETCH...... 104

LIST OF TABLES

Table page

3-1 Frequency of skeletal element number in forelimbs and hindlimbs of control and mutant animals...... 53

4-1 Summary of skeletal phenotypes among 35 Dorking incross progeny...... 76

4-2 Summary of Variability of Gene Expression, Vital Dye Staining ...... 77

B-1 Oligonucleotides used as genotyping primers...... 89

C-1 Probes used for RNA in situ hybridization...... 90

LIST OF FIGURES

Figure page

3-1 SHH and components of the hedgehog signaling pathway are present in the limb bud ectoderm ...... 39

3-2 SHH immunohistochemistry on E10.5 embryos ...... 40

3-3 Ptch1:LacZ expression in hindlimb buds ...... 41

3-4 Smo expression in the embryo including the AER of the developing limb ...... 42

3-5 Msx2-Cre is expressed robustly in the AER and not in the limb bud mesoderm ...... 43

3-6 Hedgehog signal transduction in the AER has a functional role in limb patterning ...... 44

3-7 Ptch1 is lost from the AER of Msx2-Cre;Smoflox/flox limb buds at E10.5 ...... 45

3-8 Extra condensations appear on Msx2-Cre;Smoflox/flox limbs between E13.5 and E16.5 ...... 46

3-9 Fgf8 is expressed within the morphologically distinct AER in the developing limb ...... 47

3-10 Removal of hedgehog signaling in the AER results in an increase in AER length ...... 48

3-11 Fgf4 is excluded from the posterior AER after treatment with SHH soaked beads ...... 49

3-12 Analysis of cell death in limbs that have been treated with SHH soaked beads ...... 50

3-13 Gene expression in Msx2-Cre;Smoflox/flox limbs...... 51

3-14 Model of stochastic control of AER length by hedgehog signal transduction within the AER ...... 52

4-1 Dorking chicken hindlimbs have an ectopic preaxial digit II ...... 70

4-2 Ectopic hedgehog signaling and Fgf expression occurs in the Dorking hindlimb ...... 71

4-3 Additional ectopic gene expression in Dorkings ...... 72

4-4 Ectopic Fgf4 expression in the Dorking hindlimb precedes and is independent of ectopic Shh ...... 73

4-5 Cell death is decreased in the anterior necrotic zone (ANZ) of Dorking hindlimbs...... 74

4-6 Inhibition of FGF but not Hedgehog signaling rescues cell death in the anterior necrotic zone ...... 75

A-1 β-Galactosidase activity driven by the Ptch1:LacZ allele and the Ptch1:Peak7_LacZ allele at E10.5 ...... 86

A-2 Activation of cre recombinase in tissues with both the Ptch1:Peak7CreERt2 and the R26R allele at E11.5 ...... 87

Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

MOLECULAR INTERACTIONS BETWEEN THE ZPA AND THE AER IN THE DEVELOPING VERTEBRATE LIMB

Cortney Michael Bouldin

May 2010

Chair: Brian D. Harfe Major: Medical Sciences - Genetics

The development of a vertebrate embryo is a complex process marked by several morphogenetic events, which create a highly reproducible pattern. The vertebrate limb has emerged as a model for studying pattern formation in the embryo mainly because limb manipulations do not affect embryo survival. Within the developing limb, experimental manipulation of the embryo resulted in the identification of the classical signaling centers known as the Zone of Polarizing Activity (ZPA) and the Apical

Ectodermal Ridge (AER).

The molecular signals required for function of the ZPA and AER have been identified. They are Sonic hedgehog (Shh) from the ZPA and Fibroblast Growth Factors

(Fgfs) from the AER. The functions of each of these molecules are now beginning to be understood. Analysis of Shh and hedgehog (Hh) signaling target genes has shown that

Hh activation in the limb bud mesoderm is required for normal limb development. It has

been stated that Hh signaling in the limb bud ectoderm cannot occur because

components of the Hh signaling pathway and Hh target genes have not been found in

this tissue. Contrary to previous reports, we have identified SHH protein and targets of

Hh signaling present in the limb bud ectoderm including the apex of the bud. To directly

test whether ectodermal Hh signaling was required for normal limb patterning we removed Smoothened (Smo), an essential component of the Hh signaling pathway, from the AER. Loss of functional Hh signaling in the AER resulted in disruption of the normal digit pattern and formation of additional cartilaginous condensations. These data indicate that contrary to previous accounts, the Hh signaling pathway is present and required in the developing limb AER for normal autopod development.

The formation of supernumerary digits, or polydactyly, is a common congenital

malformation. In addition to our studies investigating the role of hedghog signaling in the

AER, we characterized a spontaneous chicken mutant, known as Dorking. The

hindlimbs of Dorkings form a preaxial supernumerary digit. During the early stages of limb development ectopic expression of several genes, including Sonic Hedgehog (Shh)

and Fibroblast Growth Factor 4 (Fgf4), was found in Dorking hindlimbs. In addition to

ectopic gene expression, a decrease in cell death in the anterior of the developing

Dorking hindlimb was observed. Further molecular investigation revealed that ectopic

Fgf4 expression was initiated and maintained independent of ectopic Shh. Inhibition of

Fgf signaling but not hedgehog signaling was capable of restoring the normal anterior

domain of cell death in Dorking hindlimbs. Our data indicate that in Dorking chickens,

preaxial polydactyly is initiated independent of Shh. The sum of all findings is discussed

within the context of establishing appropriate gene expression patterns, cell number and

digit number in the developing limb.

CHAPTER 1 THE MOLECULAR BASIS OF LIMB PATTERNING

The formation of the vertebrate limb is a reproducible process that creates a complex organ. As the limb is patterned, it must be shaped along three axes. Using one’s own arm as an example, the limb forms from shoulder to digit tip along the proximal to distal axis, from the thumb to small finger along the anterior to posterior axis and from the back of the hand to the palm along the dorsal to ventral axis.

When the limb begins to grow out from the body wall, the developing limb is

composed of two cell types: loosely compacted mesenchymal cells derived from the

mesoderm and tightly associated epithelial cells derived from the ectoderm. By

removing large pieces of the developing limb, Ross Harrison observed that the

remaining mesodermal cells were capable of substituting for the removed cells

(Harrison, 1918), indicating that the cells of the limb are multipotent. Separation of

mesoderm from ectoderm followed by mixing and reshaping of the randomized

mesoderm into a bud which was then covered by ectoderm resulted in the formation of

limbs with multiple long bone and digit elements (MacCabe et al., 1973; Zwilling, 1964).

By establishing that mesodermal cells are interchangeable, these results indicate that pattern within the limb is established by cell-to-cell signaling in a process known as regulative development.

In the past two decades, a complex program of signaling molecules has been identified which regulate the positional identity of mesodermal cells within the developing limb. The combination of a large theoretical base formed by experimental embryology and recent advances in genetics and molecular biology has revealed much about cell signaling within the context of limb outgrowth and digit formation. This chapter

will outline a framework for an understanding of limb development. As limb patterning is a vast subject, special focus will be placed on two specific regions of the developing limb bud, the Apical Ectodermal Ridge (AER) and the Zone of Polarizing Activity (ZPA).

Sustained Outgrowth of the Vertebrate Limb

The Apical Ectodermal Ridge and the Progress Zone Model

After the limb mesoderm is specified, outgrowth occurs. A morphologically distinct layer of epithelium that forms at the distal most extent of the limb, known as the Apical

Ectodermal Ridge (AER), is required for limb outgrowth. First insight into the function of the AER during limb development came from experiments in which the AER was removed resulting in a truncated limb (Saunders, 1948). Early removal of the AER resulted in a more severe truncation than late removal, indicating that the time of exposure to the AER was critical in determining how much of the limb formed

(Summerbell et al., 1973).

Cells in close proximity to the AER divide rapidly (Summerbell et al., 1973). As limb growth continues, cells move further from the AER and their rate of division slows.

Because of its ability to facilitate cell division, the zone of cells in close proximity to the

AER was termed the progress zone. The progress zone model of proximal-distal axis formation states that cells in the progress zone sense the amount of time that cells spend in proximity of the AER. After growth removes cells from the influence of the

AER, cells become specified (Summerbell et al., 1973).

Specification of cells by removal from the progress zone would suggest that targeted disruption of the cells in the progress zone at intermediate stages of development should result in loss of limb structures. In support of the progress zone model of limb patterning, irradiation of cells in the progress zone results in loss of

intermediate structures (Wolpert et al., 1979). One interpretation of this result is that after irradiation the progress zone requires time to repopulate and during this time there are no cells excluded from the influence of the AER. After the progress zone has returned to its normal size, cells are once again excluded from the influence of the AER and cell specification continues.

Recent reanalysis of these experiments provided a unique interpretation of these results. Sox9, a homeobox containing transcription factor, is required for the condensation of cells into cartilage and the formation of endochondral bone (Akiyama et al., 2002; Barna and Niswander, 2007). Using Sox9 as a marker of chondrocyte specification, irradiated limbs were shown to lose cells that already expressed Sox9 and had been specified as chondrocytes (Galloway et al., 2009). According to this interpretation of irradiating limbs, the cells that are mitotically arrested by irradiation are not undifferentiated occupants of the progress zone, but cells that had already been specified to form cartilage.

After reanalysis of limb irradiation, the progress zone model still offers explanation

for limb truncations, which are observed after AER removal (Summerbell et al., 1973).

After AER removal, large amounts of cell death are observed in a region as far as 200 microns from the distal tip of the limb (Dudley et al., 2002; Rowe et al., 1982). Because the 200 microns of cells undergoing apoptosis are observed regardless of the limb stage, an alternative explanation for the limb truncation phenotypes observed after AER removal could be that cell death eliminates progenitors required to form the various segments of the limb.

A Molecular Model for Limb Outgrowth

On the molecular level, the function of the AER is to provide the limb with

Fibroblast Growth Factors (FGFs). Placement of beads soaked in FGF protein on a developing limb bud lacking an AER rescued outgrowth (Niswander et al., 1993). Four

Fgf family members are expressed in the mouse AER – Fgf4, Fgf8, Fgf9 and Fgf17

(Martin, 1998). Removal of just two of these, Fgf4 and Fgf8, limited limb bud outgrowth

(Boulet et al., 2004; Sun et al., 2002). Consistent with cell death after AER removal,

genetic removal of Fgf4 and Fgf8 resulted in cell death, but not in the distal portion of

the limb (Sun et al., 2002).

Recent studies have further clarified the role of Fgfs secreted from the AER in

proximal-distal outgrowth. In the mouse AER, four Fgfs are expressed, but they are not

expressed at the same level (Mariani et al., 2008). Because Fgfs in the AER can

functionally substitute for each another (Lu et al., 2006), removal of multiple Fgf family

members from the AER created an allelic series where developing limb buds were

exposed to a variety of FGF concentrations from the AER (Mariani et al., 2008). The

extreme example, removal of both Fgf4 and Fgf8, which are expressed at the highest

level, results in a failure of the limbs to grow out. When Fgf8, the most robustly

expressed Fgf, was reduced in combination with removal of any of the other Fgfs found

in the AER, outgrowth continued with a reduction in digit number. These results indicate

that in addition to being permissive for outgrowth, Fgfs expressed in the AER are

instructive in limb patterning (Mariani et al., 2008).

Additionally, changes in the level of FGF affect the expression of one member of a

family of proteins known as Meis genes (Mariani et al., 2008). These experiments and

others involving retinoic acid have placed the Meis genes at the center of a molecular

model for limb outgrowth. Treatment of regenerating limbs with retinoic acid induces proximal identity in regenerating tissue (Maden, 1982). Synthesis of retinoic acid occurs in the proximal portion of the limb and in the tissue adjacent to the limb, known as the flank (Mercader et al., 2000). Retinoic acid specifies limb tissue through the expression of Meis genes. When the limb is first formed, the Meis genes are expressed throughout

the limb bud mesoderm. Proliferation regulated by Fgfs at the distal extent of the limbs

separates cells from the influence of retinoic acid, silencing the expression of the Meis

genes (Mercader et al., 2000).

The developing limb can be divided into three sections. From proximal to distal,

they are the stylopod, the zeugopod and the autopod. Hoxa11 and Hoxa13 have been

used to mark the zeugopod and autopod (Yokouchi et al., 1991). Using Meis, Hoxa11

and Hoxa13 as markers of the stylopod, zeugopod and autopod respectively, a model of

proximal-distal outgrowth has been proposed that involves progressive initiation and

expansion of the domains of expression. This model relies on the spatial relationship of

cells to Fgfs from the AER and retinoic acid from the flank for establishing each of these

domains (Tabin and Wolpert, 2007).

While a molecular model of proximal-distal patterning represents a potential to

reconcile the known data relating to proximal-distal patterning and Meis genes have the

potential to be instructive in patterning (Mercader et al., 1999), Hoxa11 and Hoxa13 are

not instructive. Removal of the Hox 11 paralog group results in the production of a

normal limb (Wellik et al., 2002), and removal of Hoxa13 results in loss of only the first

digit of the limb rather than loss of the entire autopod (Fromental-Ramain et al., 1996).

Confirmation of this mechanism of proximal distal patterning will require identification of

molecules such as the Meis genes, which are instructive in the formation of the zeugopod and autopod.

Patterning within a Developing Vertebrate Limb

The Zone of Polarizing Activity (ZPA) and Molecules Expressed within the ZPA

As the limb grows, anterior to posterior patterning must be established. By looking at a fully formed forelimb, a thumb distinguishes the anterior from the posterior.

The anatomical differences between the anterior and posterior of an adult limb are controlled by a small portion of mesodermal tissue in the posterior of the limb bud known as the zone of polarizing activity (ZPA). When grafted to the anterior of the limb, the ZPA disrupts the asymmetric limb pattern and creates a limb with a mirror image duplication of the distal skeletal elements (Saunders and Gasseling, 1968).

A vertebrate homolog of the Drosophila hedgehog gene, Sonic Hedgehog (Shh), was identified and found to be expressed in the ZPA (Riddle et al., 1993). Placement of cells expressing Shh into the anterior of the developing limb reproduced the mirror image digit duplication phenotype seen upon grafting of cells located in ZPA to the anterior of the limb bud (Riddle et al., 1993). Genetic removal of Shh from the developing limb resulted in a limb with posterior defects (Chiang et al., 1996), indicating that Shh is required for the function of the ZPA and proper anterior to posterior patterning of the developing limb.

Mechanisms of Action for Shh

After the discovery of the ZPA, anterior to posterior patterning of the vertebrate digits was proposed to be established by a gradient of morphogen diffused from the

ZPA (Wolpert, 1969). Cells exposed to high concentrations of morphogen would be established as posterior and cells exposed to low concentrations would be patterned as

anterior. Consistent with the formation of a morphogen gradient, placement of increasing numbers of cells from the ZPA into the anterior of the limb created digit duplications of increasing severity (Tickle, 1981). Additionally, increasing concentrations of SHH in the anterior of a developing limb were capable of inducing mirror image duplications with increasing numbers of digits with increasingly posterior identity (Yang et al., 1997).

A morphogen model of Shh function requires that SHH be secreted to establish a gradient of SHH protein that can be interpreted by the cells of the limb field to specify different cell fates (Wolpert, 1969). Through a series of immunohistochemical analyses, a gradient of SHH protein has been identified in the posterior of the developing limb

(Gritli-Linde et al., 2001). Further, analysis of Shh expression and targets of hedgehog signaling revealed that Shh activated targets of the hedgehog signaling pathway several cell layers away from the domain of expression (Lewis et al., 2001).

To identify the fate of cells from the ZPA, cells that have expressed Shh were traced throughout limb development (Harfe et al., 2004). Descendants of Shh expressing cells were found to give rise to the posterior half of the limb. Labeling Shh expressing cells later in development revealed that late-expressing Shh descendants were restricted more posteriorly and gave rise to a smaller portion of the limb.

Comparison of cells labeled at early and late stages revealed that cells in the posterior of the limb were exposed to SHH for longer amounts of time than more anterior cells.

Since cells that give rise to the posterior most digits are exposed to SHH for the longest amount of time, a model was proposed whereby increasingly posterior digits are specified by increasing time of exposure to SHH (Harfe et al., 2004).

If the duration of SHH exposure dictates posterior digit patterning, deletion of Shh at progressively earlier time points should result in progressive loss of digits from the posterior of the limb. In mice limbs, deletion of Shh at progressively earlier time points resulted in concurrently more severe digit loss (Zhu et al., 2008). However, when the skeletons of the limbs were analyzed, the digits that were lost were found to be medial digits rather than posterior digits. Shh deletion at progressively earlier time points resulted in loss of digits in an order reverse of ossification (Zhu et al., 2008). The interpretation of this data is that only when the limb field is large enough can cells ossify to form digits. Because Shh is capable of controlling growth through modulating the expression of genes, which promote the G1-S transition (Roy and Ingham, 2002), limbs that lose Shh may no longer contain enough cells to form the appropriate number of digits. These data indicate that one of the primary roles of Shh is to regulate growth and limb field expansion.

Similar to the data uncovered in mice, the data from Towers, et. al. using the chick

model system revealed an involvement for Shh in the regulation of growth in the chick

limb. Increased Shh signaling increased expression of cyclin genes, which resulted in

increased cell division (Towers et al., 2008). However, in contrast to the results obtained

from mouse, the data from Towers et. al. reveal a continued late involvement of Shh in

patterning. By blocking cell cycle progression within the limb, Towers et. al.

demonstrated that only a single digit formed. However, the identity of the single digit

more closely resembled a digit with a posterior identity. In these experiments, Shh was

still capable of affecting pattern and promoting posterior digit identity in the absence of

cell proliferation in the limb (Towers et al., 2008).

Currently, many questions remain about the function of Shh in anterior to posterior pattering of the limb. For example, it is unclear why the studies in the chick reveal a role for Shh in both patterning and growth, while in the mouse limb Shh

appears to be mainly required for growth. Because digit identity in the mouse is more

difficult to assign than in the chick, one possibility is that the digit identity in the mouse

experiments are variable, or misassigned. This seems unlikely because several

methods were used to check digit identity (Zhu et al., 2008). A second possibility is that

there are differences in the abilities of mouse limbs and chick limbs to respond to Shh.

In Chapter 3, we identify an essential role for Shh signaling in the ectoderm of the

developing limb. A key in reconciling the function of Shh into a model will be separating

the roles of Shh in the mesoderm and the ectoderm, and characterizing the role this

signaling pathway plays in each individually.

CHAPTER 2 NETWORKS OF SIGNALING WITHIN THE DEVELOPING LIMB

Throughout development, the limb bud integrates information from multiple signaling pathways for proper patterning. This chapter will focus on the interactions between these signaling molecules and the effect of their temporal and spatial regulation on limb development.

Establishment of the AER

The initiation of limb outgrowth results from a series of epithelial-mesenchymal

interactions mediated by FGF and WNT proteins. Expression of Fgf10 in the lateral

plate mesoderm elicits a response in the overlying ectoderm (Ohuchi et al., 1997). The

ectoderm responds by activating wnt3a (Kawakami et al., 2001; Kengaku et al., 1998;

Ohuchi et al., 1997). At the boundary between the dorsal and ventral limb bud

ectoderm, the AER is formed (Altabef et al., 1997). Formation of the AER then initiates

Fgf8 expression (Kawakami et al., 2001; Kengaku et al., 1998; Ohuchi et al., 1997).

Without Fgf8 expression, Fgf10 in the mesenchyme cannot be maintained (Boulet et al.,

2004; Sun et al., 2002). Through interactions between FGF and WNT proteins, the AER

is properly positioned and maintained by Fgf10 from the underlying mesoderm.

Establishment of the ZPA

The expression of Shh in the ZPA is similarly induced through interaction between multiple signaling pathways. Retinoic acid is a known inducer of Shh expression (Riddle et al., 1993). The involvement of retinoic acid in regulating Shh expression was further confirmed by inhibition of either retinoic acid synthesis (Stratford et al., 1996) or retinoic acid reception (Helms et al., 1996), which results in loss of Shh expression. Because retinoic acid induces expression of Hox genes including Hoxb8 (Charite et al., 1994; Lu

et al., 1997), retinoic acid was proposed to induce expression of Shh with Hoxb8 acting

as an intermediate. However, genetic removal of Hox 8 paralogs does not block

expression of Shh (van den Akker et al., 2001), suggesting that some factor other than

Hoxb8 is required for Shh expression. Another molecule induced by retinoic acid is

Hand2 (Fernandez-Teran et al., 2000). Genetic removal of Hand2 results in the loss of

Shh (Charite et al., 2000). Because Hand2 is dependent on Fgf expression (Fernandez-

Teran et al., 2000), the intersection of retinoic acid, Hand2 expression and Fgf

expression initiates expression of Shh (Cohn, 2000).

Interactions between the ZPA and the AER

After the AER is induced and the ZPA initiates Shh expression, interactions

between SHH from the mesenchyme and FGFs from the AER are required to maintain

gene expression from these two signaling centers. Implantation of a source of SHH in

the anterior of the developing limb increased expression of Fgf4 in the AER (Laufer et

al., 1994), and placement of a source of FGF in the posterior of the limb (Yang and

Niswander, 1995), but not the anterior of the limb (Niswander et al., 1994) expanded

Shh expression. Additionally, genetic removal of Fgf8 reduces the expression of Shh

(Lewandoski et al., 2000) and genetic removal of Shh reduces the expression of Fgf4

(Chiang et al., 2001).

Interactions between Shh and Fgfs are mediated by Grem1. Grem1 expression is

eliminated in ld mutants (Zuniga et al., 2004). Analyis of ld mutant limbs revealed

reduced expression of Shh and Fgf4 (Haramis et al., 1995). Incorporation of Grem1 into

the interactions between SHH and Fgfs lead to the initial description of the

Shh/Grem1/Fgf feedback loop, in which SHH signals to Grem1 which then signals to

Fgfs in the AER in a positive manner.

Because Grem1 is an antagonist of Bone Morphogenetic Proteins (Bmps), Bmps are also involved in the Shh/Grem1/Fgf feedback loop. Three Bmps are expressed in the limb bud: Bmp2, Bmp4 and Bmp7 (Robert, 2007). Genetic removal of a Bmp receptor from the AER results in loss of outgrowth (Pajni-Underwood et al., 2007).

Removal of two Bmp ligands specifically from the AER of the developing limb resulted in outgrowth of the limb but defects in limb patterning (Maatouk et al., 2009). In contrast to limb buds that lack Bmp receptor in the AER, which fail to grow out, limbs missing Bmp ligands in the AER could be analyzed for changes in Grem1 expression. Consistent with a role for Bmps within the AER mediating interactions with Grem1, Grem1 expression was reduced concomitant with a reduction of Bmps in the AER (Maatouk et al., 2009).

Roles for Gene Expression Networks within the Developing Limb

Formation of new outgrowths from the embryo require the establishment of tissue within three dimensions. Interaction between multiple signaling pathways allow unique mechanisms to control pattern within the limb bud. For example, expansion of different domains of gene expression in the mouse and chick create physical boundaries for the various signaling molecules of the limb. As expression domains are separated, the

Shh/Grem1/Fgf regulatory loop breaks down in the chick limb beginning with Grem1

(Scherz et al., 2004) and in the mouse limb beginning with Fgf4 (Verheyden and Sun,

2008). From these initially reduced domains of expression the whole regulatory loop is down-regulated. Overlapping regulation through networks of signaling pathways ensures the appropriate termination of gene expression in the developing limb.

Another potential reason for regulatory networks is that these interactions exist to correct any stochastic changes in gene expression that might occur throughout development of the limb. For example, Grem1 interacts with Bmp4 (Benazet et al.,

2009). By analyzing the time required to induce changes in gene expression, Grem1 and Bmp4 interactions were identified as a separate regulatory loop within the

Shh/Grem1/Fgf feedback loop. Using multiple regulatory loops within the limb bud, variations in gene expression between limbs, individuals and environmental conditions are overcome and a limb with a normal pattern is formed (Mackem and Lewandoski,

2009)

Within this work, we present evidence for both the coordination of gene expression as a mechanism for regulating the proper patterning of the limb within three dimensions

(Chapter 3 and 4) and evidence for a regulatory loop to fine-tune levels of gene expression within the limb (Chapter 3).

CHAPTER 3 THE SHH SIGNALING PATHWAY IS PRESENT AND REQUIRED WITHIN THE VERTEBRATE LIMB BUD APICAL ECTODERMAL RIDGE FOR NORMAL AUTOPOD PATTERNING

Within the developing limb bud, exposure to SHH controls number and identity of digits produced. Proper digit pattering requires both the establishment of a concentration dependent gradient of SHH (Yang et al., 1997) and a time dependent gradient of exposure to SHH (Ahn and Joyner, 2004; Harfe et al., 2004). The ability of

SHH to pattern the forming digits is coupled to the ability of SHH to control cell proliferation (Towers et al., 2008; Zhu et al., 2008), and a full complement of digits requires tight control over the concentration and timing of Shh expression. Although

SHH is capable of limiting its own expression (Sanz-Ezquerro and Tickle, 2000), the molecular mechanism responsible for sensing the amount of SHH from the ZPA remains unknown.

Through both short-range and long-range signaling (Goetz et al., 2002), secretion of SHH protein from Shh producing cells activates a cellular signal transduction pathway. Binding of SHH protein to the transmembrane protein PTCH1 activates the Hh signaling pathway (Fuse et al., 1999). Once SHH binds to PTCH1, inhibition of another transmembrane bound protein, SMO, is relieved (Murone et al., 1999). Through a complex of proteins, SMO modulates proteolytic cleavage of members of the Ci/Gli family of transcription factors (Hooper and Scott, 2005). GLI transcription factors then directly activate transcription of hedgehog target genes, such as Gli1 and Ptch1 (Vokes et al., 2007; Vokes et al., 2008).

Within the limb, SHH is involved in a network of positive gene interactions known as the Shh/Grem1/Fgf feedback loop. SHH is capable of upregulating Fgf4 expression

in the AER (Laufer et al., 1994). Conversely, changes in FGF4 can upregulate Shh expression (Yang and Niswander, 1995). The Bmp antagonist, Gremlin1, is an intermediate for the interactions between Shh and Fgf4 (Khokha et al., 2003; Panman et al., 2006). Spatial disruption of the positive interactions of the Shh/Grem1/Fgf feedback

loop has been shown to terminate feedback loop gene expression (Scherz et al., 2004;

Verheyden and Sun, 2008). Regulation of the Shh/Grem1/Fgf feedback loop is essential

for proper patterning of the limb. Robust regulation of genes involved in the loop is

mediated by Gremlin1 and Bmp4 (Benazet et al., 2009). In addition to mutual positive interactions within the Shh/Grem1/Fgf feedback loop, the Etv4/5 genes, which are

targets of Fgf signaling, are required for proper spatial restriction of Shh in the limb bud

mesenchyme (Mao et al., 2009; Zhang et al., 2009).

Although coordinated regulation of limb development by interactions between the

ZPA and AER have been documented (Niswander, 2002), the role Hh signaling may

play within the limb bud ectoderm is unknown. This is due in part to reports that

components required for Hh signaling are not found in the ectoderm of the vertebrate

limb bud (Pearse et al., 2001; Quirk et al., 1997). Recently, the treatment of limb buds

with the teratogen acetazolamide has uncovered a potential role for Hh signaling in the

limb ectoderm. In one study, in utero exposure to acetazolamide resulted in limbs with

ectrodactyly, phenotypically similar to limbs mutant for Shh. However, Shh expression in

treated limbs was not substantially different from control limbs suggesting that loss of

digits in these animals may instead have resulted from defects in the ability of cells to

respond to SHH (Bell et al., 1999). A series of grafting experiments suggested that the

limb bud ectoderm and not the mesoderm was defective in Hh signaling in these

experiments (Bell et al., 1999). Microarray experiments in which components of the Hh signaling pathway, but not Shh, were identified in the limb bud ectoderm provided further evidence that the Hh signaling pathway may be present in the limb bud ectoderm

(Bell et al., 2005).

In this report, we demonstrate that SHH protein is present in the limb bud

ectoderm and the Hh signaling pathway is required for normal autopod patterning to

occur. Removal of Smo, an essential component of Shh signal transduction, resulted in the disruption of normal digit patterning and formation of additional cartilaginous condensations. Through analysis of mice that lacked Hh signal transduction in the AER and over-expression experiments using the chick model system, we show that the length of the AER is regulated by the Hh signaling pathway. Additionally, Hh signaling within the AER is necessary to refine the expression of genes involved in the

Shh/Grem1/Fgf feedback loop. These results uncover a novel role for Hh signal

transduction within the vertebrate limb ectoderm and provide further insight into the

molecular mechanism responsible for coordinating interactions between the ZPA and

AER.

Results

SHH Protein and Components of the Hedgehog Signaling Pathway are Present in the AER

The secreted protein SHH is produced in the mesenchyme of the limb bud ZPA.

Once produced, SHH can spread many cell diameters from its source (Goetz et al.,

2002). For the Hh signaling pathway to be active in the ectoderm of the developing limb,

SHH protein should be present in this tissue. To investigate SHH protein localization in

the limb bud ectoderm, immunohistochemistry using an antibody raised against SHH

was performed. At E10.5, SHH immunostaining was observed in both the posterior mesoderm and the posterior ectoderm of the limb bud including the AER (Fig. 3-1A-B)

(Gritli-Linde et al., 2001). SHH immunostaining was not present in the anterior mesoderm or ectoderm (Fig.3-1C).

The two additional Hh homologs present in vertebrates, Dhh and Ihh, are not

expressed concomitant with Shh in the limb bud (Bitgood and McMahon, 1995). To

confirm that the staining observed was specific for SHH protein, a section from a limb

∆/∆ bud that lacked SHH (Shh ) was analyzed in a manner identical to wild type limb

∆/∆ buds. Shh were found to lack immunostaining in both the mesoderm and the ectoderm (Fig. 3-1D). Sections through different regions of the embryo and limb bud confirmed immunoreactivity of SHH in the AER and specificity of the antibody to regions of the embryo known to express SHH (Fig. 3-2).

If Hh signaling is active within a given tissue, transcriptional targets of this pathway should be present. Active Hh signaling transcriptionally activates expression of the transcription factor Gli1 (Marigo et al., 1996a). Using a tamoxifen-inducible

Gli1CreERT2 allele (Ahn and Joyner, 2004) in combination with the cre-inducible R26R reporter allele (Soriano, 1999), we fate mapped cells that activate Gli1 in the mouse limb. Administration of tamoxifen at E9.5 resulted in detection of β-galactosidase activity in the autopod and zeugopod including ectoderm in embryos at E12.5 (Fig. 3-1E-G)

(Ahn and Joyner, 2004). To determine if the ectodermal staining observed in

Gli1CreERT2;R26R embryos required SHH, the cells activating Gli1 were analyzed in

Shh null (Shh∆/∆) embryos. In Shh null limb buds, the ectoderm was negative for β- galactosidase activity (Fig. 3-1H & I) indicating that ectodermal Gli1 expression required

activation of the Hh signaling pathway by SHH. Cells surrounding the early stage cartilaginous skeletal elements in the limb mesoderm activated the Hh signaling pathway in the absence of SHH indicating Gli1-expressing cells were present in this region of the limb (Fig. 3-1H). This staining is likely the result of IHH activation of the Hh signaling pathway since Ihh is expressed in chondrocytes (Bitgood and McMahon,

1995; St-Jacques et al., 1999). These data indicate that ectodermal cells have activated the Hh signaling pathway in response to SHH produced from the ZPA.

To confirm that Hh signaling is present in the limb bud ectoderm we analyzed a second Hh pathway target gene, Ptch1 (Marigo et al., 1996b). Mice with a LacZ cassette inserted downstream of the endogenous Ptch1 promoter produce β- galactosidase activity in cells that have activated transcription from the Ptch1 locus

(Goodrich et al., 1997). In the limbs of embryos containing the Ptch1:LacZ allele, β- galactosidase activity was detected at E11.5 in the posterior limb bud ectoderm including the posterior AER (Fig. 3-1J & K). Further analysis revealed that β- galactosidase activity initially appeared punctate in the posterior of the limb including the ectoderm at E10.0 with robust expression in the ectoderm observed from E10.5 through E12.5 (Fig. 3-3). Sectioned limbs contained β-galactosidase activity in both the

posterior mesoderm and ectoderm (Fig. 3-1L). Examination of sections revealed

detectable β-galactosidase activity in the posterior AER, the posterior dorsal and ventral

ectoderm, and the ectoderm of the posterior margin (the space from the posterior AER

to the flank; Fig. 3-1L and 3-3). These data indicate that SHH is present and capable of

activating the Hh signaling cascade within the limb bud ectoderm.

Hedgehog Signaling in the AER is Required for Formation of a Normal Autopod

Although Hh signal transduction was found in several regions of the ectoderm, we chose to focus specifically on activation of the Hh signaling pathway in the AER because of the importance of the AER in limb development. SMO, a transmembrane protein required for active Hh signaling (Zhang et al., 2001), was detected in the limb bud ectoderm at E11.5 (Fig. 3-4). The Msx2-Cre allele expresses cre recombinase in

the AER but not in the limb mesoderm (Fig. 3-5) (Barrow et al., 2003). o determine if

Hh signaling in the AER played a functional role in limb patterning, we removed Smo

from the AER using Msx2-Cre and a conditional allele for Smo (see Experimental

Procedures for details). Embryos that were homozygous for the Smoflox allele and contained the Msx2-Cre transgene have lost SMO from the AER. As a result of SMO loss, Hh signaling was absent.

Loss of Hh signaling in the ectoderm of embryos in which Smo was removed from the AER was confirmed by performing Ptch1 in situ hybridizations using BM purple as a

substrate (see Experimental Procedures). Whole mount RNA in situ hybridizations to detect Ptch1 at E11.5 revealed expression in the posterior AER of control limbs but not in the AER of Msx2-Cre;Smoflox/flox limbs at the same somite stage (ss) (Fig. 3-6A-D).

Sectioned tissue revealed that in Msx2-Cre;Smoflox/flox embryos at E10.5 (Fig. 3-7) and

E11.5 (Fig 3-6E & F) Ptch1 mRNA was absent from the AER and ventral ectoderm.

Ptch1 was still expressed in the dorsal ectoderm in addition to the ectoderm proximal to the posterior AER. In situ hybridization with Ptch1 riboprobe indicated that Msx2-

Cre;Smoflox/flox limbs lacked the ability to activate the Hh signaling pathway within the

AER.

To determine whether Hh signaling within the AER played a functional role in the establishment of limb pattern, Msx2-Cre;Smoflox/flox and wild type skeletons were examined. Skeletal analysis revealed that the stylopodal and zeugopodal elements of both Msx2-Cre;Smoflox/flox and control limbs were normal. The autopods of the hindlimbs

and forelimbs of Msx2-Cre;Smoflox/flox animals exhibited extra cartilaginous condensations (Fig. 3-6G-N), which were highly penetrant (Table 3-1). Extra cartilaginous condensations were investigated using both Sox9 expression and bright field microscopy, and were found to be formed between E13.5 and E16.5 (Fig. 3-8).

These data indicate that Hh signaling within the AER is necessary for the formation of a properly patterned limb autopod. Msx2-Cre;Smoflox/flox mice die at birth, which limited our

ability to perform a detailed analysis of extra digit condensations.

AER Length is Controlled by Hh Signaling within the AER

Signals from the ZPA have been postulated to control the length and shape of the

AER (Lee and Tickle, 1985; Todt and Fallon, 1984). To investigate if Hh signaling within the limb bud ectoderm regulated the length of the AER, Fgf8 expression was examined and compared in Msx2-Cre;Smoflox/flox and control limbs. Fgf8 is expressed

throughout the entire AER and can be used to measure the length of this structure (Fig.

3-9) (Crossley and Martin, 1995; Mahmood et al., 1995). The Straighten plug-in for NIH

ImageJ was used to convert the curved AER into a straight-line, which was then

measured (see Methods). Somite matched Msx2-Cre;Smoflox/flox forelimbs (50ss, n=9,

Fig. 3-10B) and hindlimbs (50ss, n=10, Fig. 3-10D), and control forelimbs (50ss, n=4,

Fig. 3-10A) and hindlimbs (50ss, n=7, Fig. 3-10C) were collected and analyzed for Fgf8

expression. Stained AERs were then straightened and measured (Fig. 3-10E). AERs in

which Hh signaling was removed were found to be on average 700 microns (forelimbs)

and 260 microns (hindlimbs) longer than normal limbs at the same somite stage. The means were compared using a t-test (Welch’s, unpaired), which revealed the increase

in AER length in the Msx2-Cre;Smoflox/flox limbs to be statistically significant (forelimbs, p

= 0.03 and hindlimbs, p < 0.0001).

Our experiments using the mouse model system indicated that loss of Hh signaling from the AER resulted in an increase in AER length. Using the chick model system, we investigated the effects of elevated amounts of Hh protein on AER length. A

SHH-soaked bead placed just anterior to the normal ZPA was used to expand the domain of SHH protein (Fig. 3-10F). After 24 hours of treatment with a SHH-soaked bead, the length of the AER was decreased compared to the untreated contralateral limb buds or PBS treated limb buds as measured by Fgf8 expression (3 of 4 treated limbs; Fig. 3-10G-J) and Fgf4 expression (Fig. 3-11). Consistent with published reports

(Sanz-Ezquerro and Tickle, 2000), placement of a SHH-soaked bead in the posterior limb bud mesenchyme resulted in an increase in cell death in both the AER and the mesoderm of the limb bud (Fig. 3-12) but did not cause defects in the patterning of limb skeletal elements indicating that the limb bud recovers from transient exposure to elevated levels of SHH (data not shown).

Hedgehog Signaling in the AER Regulates the Shh/Grem1/Fgf Feedback Loop

The expression of the genes involved in the Shh/Grem1/Fgf feedback loop is dynamic and dependent on the timing and levels of other components in the regulatory network (Benazet et al., 2009; Panman et al., 2006; Scherz et al., 2004; Verheyden and

Sun, 2008). To investigate the effect of Hh signaling within the AER on the components of the Shh/Grem1/Fgf feedback loop, double RNA in situ hybridizations for Shh and

Fgf4 or Fgf8 were performed on normal and Msx2-Cre;Smoflox/flox embryos. Fgf4

expression was elevated in Msx2-Cre;Smoflox/flox limbs compared to wild type limbs (Fig.

3-13A & B). Expression of Fgf8, which interacts with Fgf4 (Lewandoski et al., 2000), was prolonged in Msx2-Cre;Smoflox/flox limbs (Fig. 3-13G & H).

Consistent with a link between Fgfs and the expression of Shh (Lewandoski et al.,

2000; Sun et al., 2000), loss of Hh signaling in the AER resulted in increased

expression of Shh in the underlying mesenchyme for longer amounts of time than in

somite matched control limbs (Fig. 3-13A-D). Analysis of Msx2-Cre;Smoflox/flox limbs for

Gremlin1 expression revealed that the domain of expression was more broad when compared to somite matched control limbs (Fig. 3-13E & F). These data indicate that proper regulation of the Shh/Grem1/Fgf feedback loop requires activation of the Hh signaling pathway in the limb bud AER.

Discussion

Within the vertebrate limb mesoderm, several different mechanisms of patterning have been reported for SHH. SHH acts through a concentration gradient (Yang et al.,

1997) and a temporal gradient (Harfe et al., 2004) to establish both digit identity and number. Additionally, through regulation of genes that promote the G1-S transition, SHH affects limb patterning by controlling growth (Towers et al., 2008; Zhu et al., 2008). In this study, we present evidence that Hh signaling is present in the AER and is required within the AER for correct patterning of the vertebrate autopod.

Activation of the Hh signaling pathway was detected in several domains within the posterior ectoderm of the developing limb. Because Shh expression has not been detected by RNA in situ hybridization (Riddle et al., 1993) nor microarrays (Bell et al.,

2005) in the limb bud ectoderm, Hh signals to this tissue through a paracrine

mechanism. Targets of the Hh signaling pathway were found in the AER, the dorsal and

ventral ectoderm, and the ectoderm of the posterior margin (ectoderm between the AER and the flank). After removal of Hh signaling from the AER, Hh signaling remained in ectodermal cells juxtaposed to the posterior AER. The lack of a stronger phenotype upon removal of Hh signaling from the AER may be due to the presence of Hh responding cells in the posterior margin. Active hedgehog signaling in this region of the limb bud may be able to compensate for a loss of Hh signaling in the AER. Removal of

Hh signaling from both the AER and the posterior margin has the potential to result in a more significant expansion of the AER and an increase in the severity of polydactyly.

Upon removal of Hh signaling from the AER, we observed changes in gene expression of components of the Shh/Grem1/Fgf feedback loop. Previously, Shh expression in the ZPA was found to be regulated by Fgfs expressed in the AER

(Lewandoski et al., 2000; Lu et al., 2006). In addition, Gremlin1 expression in the mesoderm is controlled by SHH (Nissim et al., 2006; Vokes et al., 2008). In our experiments, removal of Hh signaling from the AER increased the length of the AER, resulting in an increase in the number of cells capable of expressing Fgfs. Because recombinase expression was specific to the ectoderm, the observed alterations in components of the Shh/Grem1/Fgf feedback loop are predicted to result from an increase in the length of the AER caused by a loss of Hh signal transduction within this tissue.

The amount of SHH produced in the limb bud has been shown to control the number of Shh expressing cells in the ZPA through cell death (Sanz-Ezquerro and

Tickle, 2000). Consistent with the observations of Sanz-Ezquerro et. al., our

experiments increasing SHH in the posterior of the chick limb induced cell death. In

addition to an increase in cell death, we identified a reduction in AER length, detected by Fgf8 and Fgf4, expression after elevating SHH protein levels in the limb bud mesenchyme. Our experiments, in which we either genetically removed Hh signaling from the AER or ectopically elevated SHH protein in the limb mesenchyme, indicate that

Hh signaling in the AER refines the length of the AER. We propose that changes to the

AER in response to the Hh signaling pathway during normal limb development refine the initial levels of SHH produced by the ZPA through changes induced by Fgf

expression.

During normal limb development, the initial levels of SHH produced by the ZPA is

likely to vary between embryos and possibly even within the limbs buds of a given

embryo. To ensure normal patterning, the pool of Shh expressing cells has to be refined

to produce an exact and reproducible amount of SHH in every limb bud. We propose

that during normal development, if the original pool of cells in the ZPA produces too

much SHH, there is a corresponding increase in Hh signaling in the AER resulting in a

decrease in AER length. SHH producing cells are then removed from the ZPA (Sanz-

Ezquerro and Tickle, 2000). If too little SHH is produced, Hh signaling in the AER

decreases, resulting in an increase in AER length and a corresponding increase in SHH

production in the ZPA (see model in Fig. 3-14). In our model, the ability of the AER to

directly respond to SHH facilitates the production of a reproducible concentration of

SHH in every limb bud within a given species. In normal limbs, the proposed changes in

the length of the AER and SHH produced from the ZPA are most likely very subtle since

large alterations in either SHH protein or AER length would result in defects in autopod

patterning.

Regulation of Shh expression in the mesoderm of fins and appendages has been identified as a potential source of morphological change over the course of evolution

(Dahn et al., 2007). We propose that the response to Hh signaling within the ectoderm could also function as a source of evolutionary change. Since Hh signaling in the ectoderm is required for patterning the autopod, it is possible that by changing the ability of limb bud ectodermal cells to respond to SHH, digit number could be altered during evolution.

Experimental Procedures

Mouse Alleles and Breeding

The Shh∆, Gli1-CreERT2, R26R, Ptch1:LacZ, Msx2-Cre and Smoflox alleles were maintained and genotyped as previously described (Ahn and Joyner, 2004; Chiang et al., 1996; Goodrich et al., 1997; Soriano, 1999; Sun et al., 2000; Zhang et al., 2001).

Genotyping was performed on DNA isolated from tail or yolk sack tissue. Embryos harvested at E12.5 or earlier were staged by counting somites. For experiments involving embryos lacking Smo in the AER, control embryos lacked the Msx2-Cre allele

and were either heterozygous or homozygous for the Smoflox allele. Both genetic

combinations were phenotypically indistinguishable from normal mice. Alleles were

maintained on mixed backgrounds, and animals were handled in accordance with the

University of Florida IACUC.

Fate Mapping, β-Galactosidase Detection and Immunohistochemistry

Tamoxifen was administered in corn oil (Sigma C-8267) at a final concentration

of 20 mg/ml by oral gavage as previously described (Ahn and Joyner, 2004). For whole

mount detection of β-galactosidase activity, embryos were harvested and fixed

overnight in 0.2% formaldehyde. After fixation, β-galactosidase activity was detected as previously described (Harfe et al., 2004). For β-galactosidase detection on sections,

embryos were fixed and cryoprotected with 30% Sucrose in PBS. After embedding in

O.C.T. compound (Sakura 4583), cryoprotected embryos were sectioned at 12 microns.

Slide mounted tissue was washed two times with PBS before being incubated overnight

at 37°C in staining solution [1 mg/ml X-gal, 5 mM K3Fe(CN)6 and 5 mM K4Fe(CN)6 in

concentrated rinse buffer (0.1 M NaH2PO4, pH 7.4, 2 mM MgCl2, 0.2% Igepal, 0.1%

sodium deoxycholate)]. Stained tissue was rinsed three times with PBS, dehydrated

and mounted with Permount (Fisher SP15-500). SHH immunohistochemistry was

performed as previously described (Gritli-Linde et al., 2001).

Whole Mount RNA in situ Hybridization and AER Measurement

Embryos harvested for in situ hybridization were incubated overnight in 4%

formaldehyde at 4°C. Section and whole mount RNA in situ hybridizations were

performed according to a standard protocol (Wilkinson and Nieto, 1993) with the

exception that BM Purple (Roche, 1442074) was used in place of NBT and BCIP to

develop Ptch1 in situs. Smo riboprobe (used in Fig. 3-4) synthesis was performed on a

plasmid constructed by RT-PCR using the following primers: forward, 5’ –

ttcccagggttgaagacag, reverse, 5’ – cacgttgtagcgcaaag. AER length was calculated by

importing pictures into NIH ImageJ (Rasband, 1997-2007). A calibration slide was

measured at the same magnification to set the scale. The “Straighten” plug-in was used

to turn the curved AERs into measurable straight-lines (Kocsis et al., 1991). The mean

AER length of Msx2-Cre;Smoflox/flox forelimbs (50ss, n=9) and hindlimbs (50ss, n=10),

and control forelimbs (50ss, n=4) and hindlimbs (50ss, n=7) was collected, and a t-test

(Welch’s, unpaired) was used to test the significance (α = 0.05).

Bead Implantation and LysoTracker Analysis

Affi-gel beads (Bio Rad, 153-7302) were soaked in hrSHH (1mg/mL in PBS; R &

D Systems, 1845-SH) or PBS for at least one hour. Chicks were staged according to the staging guide by Hamburger and Hamilton (Hamburger and Hamilton, 1951). Eggs were fenestrated with laminectomy forceps. At HH stage 20, the amnionic raphe was separated and a bead was transferred to the egg using watchmaker’s forceps. Next, a microdissection needle was used to create a small hole in the mesenchyme proximal to the AER in a region just anterior to the ZPA. The bead was then inserted into the space created by the microdissecting needle, and the embryos were placed at 37°C overnight.

Embryos were harvested after 24 hours of incubation and fixed in 4% formaldehyde at

4°C overnight before further analysis. LysoTracker (Molecular Probes, L-7528), an acidophilic dye which accumulates in the lysosmes of apoptotic cells and phagocytic cells engulfing apoptotic cells (Zucker et al., 1999), was used as previously described

(Bouldin and Harfe, 2009).

Figure 3-1. SHH and components of the hedgehog signaling pathway are present in the limb bud ectoderm. Shh immunohistochemistry. A) At low magnification, SHH is detected in a graded fashion specifically in the posterior of the limb. B) At high magnification, SHH was visible in the posterior limb bud ectoderm including the AER. C) SHH was not visible in the anterior limb bud ectoderm. In B & C, the arrows highlight the AER, and the white dashed lines separate the mesoderm from the ectoderm. D) Limb bud tissue from a Shh∆/∆ limb, which was mounted on the same slide as the tissue shown in images A-C, had no detectable SHH. Fate-mapping of cells responding to Hh signal transduction in limbs exposed to tamoxifen at E9.5 and harvested at E12.5. Sectioned limbs of Gli1-CreERT2;R26R embryos showed descendants of cells responding to Hh signal transduction in the posterior mesoderm and posterior ectoderm of the E) autopod and zeugopod F) at low magnification and G) high magnification. Sectioned Shh∆/∆ limbs containing the Gli1-CreERT2 and R26R alleles contained no cells that had responded to Hh signaling in the ectoderm of the zeugopod H) at low magnification and I) high magnification. β-galactosidase activity in limbs containing the Ptch1:LacZ allele. Whole mount hindlimbs at E11.5 showed active Hh signaling in the posterior limb mesoderm and ectoderm including the AER J) at low magnification and K) high magnification. L) Sectioned hindlimbs confirmed the presence of β- galactosidase activity in the posterior ectoderm (arrowhead denotes staining in the ectoderm). The plane of section of L is illustrated by a black line in K. Images in A-D provided by Amel-Gritli Linde and images in E-I provided by Sohyun Ahn.

Figure 3-2. SHH immunohistochemistry on E10.5 embryos. To confirm the ectodermal immunoreactivity of SHH and the specificity of the antibody, limb buds were analyzed from different planes of section. Embryos at E10.5 cut at the transverse plane of section showed immunoreactivity of SHH in several tissues known to be positive for SHH including the gut (G), the notochord (N), the neural tube (NT) and the posterior region of the limb including the ectoderm A) at low magnification and B) high magnification. Similarly, embryos at E10.5 cut on an oblique plane of section showed the immunoreactivity of SHH in tissues known to express Shh and in the posterior region of the limb including the ectoderm C) at low magnification and D) high magnification. The box in C is enlarged in D. The arrow in D highlights SHH immunostaining in the AER. Images provided courtesy of Amel Gritli-Linde.

Figure 3-3. Ptch1:LacZ expression in hindlimb buds. To expand our analysis of Ptch1 expression in the limb bud ectoderm, we analyzed Ptch1:LacZ embryos at various stages. A) At E10.0, β-Galactosidase activity was punctate in the posterior of the developing limb at low magnification. B) At a higher magnification, β-Galactosidase activity was present in the posterior of the developing limb including the ectoderm (highlighted by an arrowhead). At low magnification C) in E10.5 and E) E11.5 llimbs, β-Galactosidase activity was present in the posterior of the limb. At high magnification in D) E10.5 and F) E11.5 robust staining was visible in the ectoderm (highlighted by an arrowhead). G) At E12.5, β-Galactosidase activity was no longer posteriorly restricted in the mesoderm and was found surrounding proliferating chondrocytes, a known source of Ihh. H) At high magnification in the E12.5 limb, β-Galactosidase activity persisted in the posterior ectoderm (highlighted by arrowheads). Additional sagittal sections from embryos containing the Ptch1:LacZ allele at E11.5 were analyzed in detail on sections. I) A schematic of the posterior of the developing limb bud showing approximate locations of sectioned tissue. J) In a section from the posterior margin (the posterior region from the flank to the AER), β-galactosidase activity was detectable in the posterior ectoderm including portions of the dorsal and ventral ectoderm. K) In a section from the proximal portion of the posterior AER, β- galactosidase activity was detectable in the ectoderm including the AER. L) In a section from a more distal portion of the posterior AER revealed β- galactosidase activity exclusively in the AER. Arrows mark the dorsal and ventral extremes of detectable β-galactosidase activity in the ectoderm.

Figure 3-4. Smo expression in the embryo including the AER of the developing limb. To investigate the presence of Smo in the embryo, a series of RNA in situ hybridizations were performed using a riboprobe designed to detect Smo. A) At 37ss, embryos hybridized to a sense probe revealed no staining, however, B) embryos stained with an antisense probe expressed Smo throughout the embryo including C) in the limb. Tissue harvested at 50ss and sectioned revealed staining throughout the mesoderm and ectoderm of the limb including the AER D) at low magnification and E) high magnification. The arrow in E highlights Smo expression in the AER.

Figure 3-5. Msx2-Cre is expressed robustly in the AER and not in the limb bud mesoderm. To investigate the expression of Msx2-Cre in the developing limb, mating schemes were designed to incorporate the Rosa26 reporter into animals with the Msx2-Cre allele. In these limbs, cells that have β- galactosidase activity have at some time during development expressed cre recombinase from the Msx2-Cre allele. At E10.5, Msx2-Cre was expressed in the AER of both A) forelimbs and B) hindlimbs. C & D) Sagittal sections of E10.5 limbs revealed β-galactosidase activity in the AER and ventral ectoderm of two different limbs. In all of the limbs analyzed, β-galactosidase activity was restricted to the ectoderm.

Figure 3-6. Hedgehog signal transduction in the AER has a functional role in limb patterning. RNA in situ hybridizations with control and Msx2-Cre;Smoflox/flox limbs using a riboprobe for Ptch1. Ptch1 was expressed throughout the posterior of the control limb at 47ss including the ectoderm A) at low magnification and C) high magnification. In Msx2-Cre;Smoflox/flox limbs at 47ss, Ptch1 is expressed in the posterior limb bud mesoderm and in the most proximal region of the posterior ectoderm, but is not expressed in the AER B) at low magnification and C) high magnification (arrowhead denotes loss of expression in the AER while the arrow marks expression in ectoderm adjacent to the AER). F) Sectioned limbs revealed that Ptch1 was absent from the AER of Msx2-Cre;Smoflox/flox limbs but E) present in all posterior ectoderm of control limbs (arrowhead denotes loss of expression in the AER while the arrow marks expression in ectoderm adjacent to the AER). In C-F, the white dashed lines highlight the visible boundaries of the mesoderm and ectoderm. In C & D, the solid white lines highlight the posterior edge of the limb. Skeletal analysis of P0 limbs of control forelimbs G) at low magnification and I) high magnification and mutant forelimbs H) at low magnifcation and J) high magnification. Skeletal analysis of P0 limbs of control hindlimbs K) at low magnification and M) high magnification and mutant forelimbs L) at low magnifcation and N) high magnification. and hindlimbs Insets in J and N show the supernumerary cartilaginous condensations found in Msx2-Cre;Smoflox/flox animals. s = scapula, ss = spine of the scapula, h = humerus, dt = deltoid tuberosity, r = radius, u = ulna, a = autopod, cb = carpal bones, mc = metacarpals, il = ilium, is = ischium, f = femur, p = patella, t = tibia, fi = fibula, tb = tarsal bones, mt = metatarsals.

Figure 3-7. Ptch1 is lost from the AER of Msx2-Cre;Smoflox/flox limb buds at E10.5. To expand our analysis of the loss of Shh signaling in the developing limb ectoderm, we analyzed expression of Ptch1 by whole mount RNA in situ hybridization. Ptch1 was expressed throughout the posterior of the control limb at 35ss including the ectoderm A) by whole mount and C) section. B) In Msx2-Cre;Smoflox/flox limbs at 35ss, Ptch1 was expressed in the posterior limb bud mesoderm and in the most proximal region of the posterior ectoderm, but was not expressed in the AER. Arrowhead denotes loss of expression in the AER while the arrow marks expression in ectoderm adjacent to the AER. C) Sectioned limbs revealed that Ptch1 was present in posterior ectoderm of control limbs, D) but absent from the AER of Msx2-Cre;Smoflox/flox limbs (arrowhead denotes loss of expression in the AER while the arrow marks expression in ectoderm adjacent to the AER).

Figure 3-8. Extra condensations appear on Msx2-Cre;Smoflox/flox limbs between E13.5 and E16.5. To further investigate the extra cartilaginous condensations formed in Msx2-Cre;Smoflox/flox limbs, limbs were analyzed at E13.5 for expression of Sox9 and at E16.5 by bright field microcopy. Sox9 expression A) in control forelimbs and B) Msx2-Cre;Smoflox/flox forelimbs and C) control hindlimbs and D) Msx2-Cre;Smoflox/flox hindlimbs by whole mount RNA in situ hybridization were similar. However, analysis of A) control forelimbs and B) Msx2-Cre;Smoflox/flox forelimbs and C) control hindlimbs and D) Msx2- Cre;Smoflox/flox hindlimbs forelimbs (E & F) and hindlimbs (G & H) by bright field microscopy revealed an extra cartilaginous condensation at E16.5 in the posterior of Msx2-Cre;Smoflox/flox limbs.

Figure 3-9. Fgf8 is expressed within the morphologically distinct AER in the developing limb. To confirm previous reports that Fgf8 was expressed throughout the entire AER, limbs were analyzed by RNA in situ hybridization for expression of Fgf8. A) Analysis of Fgf8 expression revealed robust expression throughout the AER in 43ss hindlimbs. B) Analysis of Fgf8 expression on serial sections from embryos at 39ss revealed that when the ectoderm was a simple epithelial layer, no Fgf8 was expressed. C & D) Fgf8 was expressed when the epithelium was stratified. White dashed line denotes the boundary between mesenchyme and epithelium. B-D were taken at the same magnification. Scale bar = 10 μm

Figure 3-10. Removal of hedgehog signaling in the AER results in an increase in AER length. Double RNA in situ hybridizations on A) control forelimbs, B) Msx2- Cre;Smoflox/flox forelimbs,C) control hindlimbs and D) Msx2-Cre;Smoflox/flox hindlimbs using a riboprobe against Fgf8 and Shh at 50ss. Both forelimbs and hindlimbs. AERs were larger in Msx2-Cre;Smoflox/flox embryos than in somite- matched controls. E) Average AER length after measurement of Fgf8 expression in Msx2-Cre;Smoflox/flox limbs and control limbs using the Straighten plug-in for NIH ImageJ. Data are represented as means and the error bars represent the SEM. A t-test (Welch’s, unpaired) indicated that these differences were statistically significant (* = p = 0.03 and ‡ = p < 0.0001). F) A diagram of chick limbs with a bead soaked in SHH implanted just anterior to the ZPA in the distal limb bud mesenchyme. G) Fgf8 expression was decreased after 24 hours of treatment with a SHH soaked bead beginning at HH 20, H) when compared to an untreated contralateral control limb. The arrow in C highlights the loss of posterior AER in the treated limb bud. I) Fgf8 expression after 24 hours of treatment with a PBS soaked bead was unchanged, J) when compared to a contralateral control limb. The proximal location of the beads 24 hours after implantation is a result of outgrowth of the limb bud.

Figure 3-11. Fgf4 is excluded from the posterior AER after treatment with SHH soaked beads. To determine if increased SHH protein in the posterior of the limb altered the domain of Fgf4 expression, we treated limbs with beads soaked in SHH (see Experimental Procedures for more detail). A) After 24 hours of treatment with a SHH soaked bead, Fgf4 expression was reduced in the posterior AER, B) when compared to the contralateral control. C) PBS soaked beads had no effect on the expression of Fgf4, D) when compared to the contralateral control.

Figure 3-12. Analysis of cell death in limbs that have been treated with SHH soaked beads. To determine if increased SHH protein in the posterior of the limb induced changes in cell death, we analyzed cell death using LysoTracker red in limbs treated with a SHH soaked bead (see Experimental Procedures for more detail). A) Analysis of cell death in limbs after treatment with PBS or C) in an untreated contralateral control revealed cell death in the anterior necrotic zone and the AER. B) After 24 hours of treatment with a SHH soaked bead, cell death was noticeably increased in the posterior of the limb including the mesoderm and AER (9/13 limbs). In A & B, white circles depict bead placement. The black arrow in B highlights cell death in the posterior of the limb bud and the white arrowhead highlights cell death in the posterior AER. To determine the effect of the SHH bead on the Fgf8 expression, we analyzed Fgf8 expression in the same limb buds used in A-C. D) Fgf8 expression was unaffected by treatment with a PBS soaked bead or F) in a contralateral control limb. E) Treatment with a SHH soaked bead reduced the domain of Fgf8 expression in the developing limb bud. Merged images reveal that G) in a PBS treated and I) a contralateral control limb, cell death can be detected at the anterior and posterior extremes of the domain of Fgf8 expression. H) Treatment with a SHH soaked bead resulted in a noticeable increase in cell death with much of the observed cell death occurring at the anterior and posterior extremes of the domain of Fgf8 expression.

Figure 3-13. Gene expression in Msx2-Cre;Smoflox/flox limbs. Double labeled RNA in situ hybridizations of A) control and B) Msx2-Cre;Smoflox/flox hindlimbs for Shh (red) and Fgf4 (purple), revealed an increase in Fgf4 and Shh expression at 39ss. Analysis of Fgf8 expression in C) 61ss control hindlimbs and D) 63ss Msx2-Cre;Smoflox/flox hindlimbs revealed that Fgf8 expression was prolonged in Msx2-Cre;Smoflox/flox compared to control limbs. Arrows in D denote prolonged expression of Fgf8. Expression of Gremlin was expanded at 50ss in F) Msx2-Cre;Smoflox/flox compared to E) control hindlimbs. White lines show the width of the domain of Gremlin expression.

Figure 3-14. Model of stochastic control of AER length by hedgehog signal transduction within the AER. A) During normal development a population of cells within the ZPA expresses Shh. Response to Hh signaling within the ectoderm limits or expands the length of the AER. B) If levels of SHH protein in the AER are elevated, the AER is shortened due to elevated Hh signaling in this tissue. Shortening of the AER results in a corresponding decrease in mesenchymal Shh expression. C) If levels of SHH protein is decreased or absent from the AER, decreased Hh signaling occurs in the AER and the AER expands in length. As a result of a longer AER, there is a corresponding increase in Shh expression in the underlying mesenchyme. This buffering model ensures that the amount of SHH produced from the ZPA is appropriate and results in a reproducible digit pattern. In A-C only the posterior of the limb bud is diagramed. Green in the mesenchyme shows the level of SHH produced in the ZPA, while red in the AER denotes Hh signaling. Dark blue denotes the posterior AER, where we identified targets of Hh signaling. The more anterior AER is show as light blue.

Table 3-1. Frequency of skeletal element number in forelimbs and hindlimbs of control and mutant animals. # of Digits in the Autopod Genotype Limbs n Observed (percent) Less Six Five than five 18 Forelimb Control 18 0 0 (100%) Msx2- flox/flox 8 2 Cre;Smo 10 0 (80%) (20%) * 18 Hindlimb Control 18 0 0 (100%) Msx2- 9 1 Cre;Smoflox/flox 10 0 (90%) (10%) * * Samples were analyzed immediately after fixation and before skeletal preparation due to potential loss of extra tissue during the clearing process.

CHAPTER 4 ABERRANT FGF SIGNALING, INDEPENDENT OF ECTOPIC HEDGEHOG SIGNALING, INITIATES PREAXIAL POLYDACTYLY IN DORKING CHICKENS

Introduction

Polydactyly, the most frequent congenital hand malformation observed in humans, has a prevalence between 5 and 19 per 10,000 live births, (Sesgin and Stark, 1961;

Zguricas et al., 1999). Over one-hundred clinically distinct disorders have polydactyly as a symptom and, approximately two-thirds have an unidentified molecular cause

(Biesecker, 2002). A more complete understanding of polydactyly relies on a better understanding of limb development and the etiology of this disorder.

Outgrowth and the initial steps of limb patterning are controlled by at least two classically described signaling centers: the zone of polarizing activity (ZPA) and the apical ectodermal ridge (AER) (Saunders, 1948; Saunders and Gasseling, 1968). The

ZPA is a posterior portion of mesenchyme, which when transplanted to the anterior of a host limb produces mirror image duplications (Saunders and Gasseling, 1968). The molecule responsible for mediating ZPA activity has been identified as SHH. Further, implantation of a source of SHH protein in the anterior of a developing limb bud induces the formation of preaxial extra digits (Riddle et al., 1993).

The AER is a distal portion of the ectoderm, which is required for limb outgrowth

(Saunders, 1948). One of the primary functions of the AER is to express Fibroblast growth factors (FGFs) [reviewed in (Martin, 1998)]. The requirement for AER-FGFs has been demonstrated by placing FGF soaked beads on the distal end of limbs in which the AER had been removed. In these experiments, FGF protein sustained limb outgrowth (Fallon et al., 1994; Niswander et al., 1993). Genetic removal of two of the four Fgfs expressed in the mouse AER results in limb truncation (Boulet et al., 2004;

Sun et al., 2002). Recent studies indicate that the AER-FGFs are functionally equivalent, and differences in expression level, timing of expression and receptor affinity make the AER-FGFs unique during limb patterning (Lu et al., 2006; Mariani et al.,

2008)

Identification and characterization of the molecules required for ZPA and AER function have indicated that these genes are involved in a feedback loop. For example, expression of three of the four ectodermally expressed Fgfs in mouse require Shh for their maintenance (Chiang et al., 1996; Sun et al., 2000), while removal of multiple Fgfs

from the AER results in a loss of Shh (Mariani et al., 2008; Sun et al., 2002).

Interestingly, SHH is capable of inducing expression of Fgf4. For example, placement

of a source of SHH in the anterior of the limb bud causes Fgf4 to expand anteriorly in

the AER (Laufer et al., 1994; Niswander et al., 1994). The coordinated regulation of

gene expression between the AER and the underlying mesenchyme provides a

mechanism for integrating patterning along multiple axes of development (Niswander,

2002). Also, coordinate regulation provides a mechanism for the termination of gene

expression (Scherz et al., 2004; Verheyden and Sun, 2008).

Changes in expression of Shh and Fgfs can be used to describe the etiology of

many of the characterized cases of polydactyly. Several mutants with preaxial

polydactyly have been identified in which anterior expression of Shh occurred.

Additionally, these mutants with preaxial polydactyly have been shown to have Fgf4

ectopically expanded anteriorly in the AER (Chan et al., 1995; Masuya et al., 1997;

Masuya et al., 1995; Sharpe et al., 1999). Genetic analysis has revealed that two of

these mutants, Hemimelic extra toes and Sasquatch, had mutations in an enhancer

element upstream of the Shh locus resulting in ectopic anterior expression of Shh

(Lettice et al., 2003; Lettice et al., 2002).

The goal of the present study was to use the Dorking breed of chickens to identify molecular pathways responsible for polydactyly in vertebrates. Differences in Shh and

Fgf expression in addition to cell death were found in Dorking hindlimbs when compared to wild type. By taking advantage of the in ovo development of chickens, we uncovered a molecular mechanism in which polydactyly was initiated independently of ectopic Shh.

Results

Dorking Chickens have Partially Penetrant Preaxial Polydactyly in the Hindlimb but not in the Forelimb.

Polydactyly in Dorkings has been previously reported to exhibit a dominant mode of inheritance with variable penetrance (Punnett and Pease, 1929). An outcross of our population confirmed the dominance of the Dorking phenotype with 92% (n=12) of offspring showing polydactyly. To characterize the hindlimbs of Dorkings (Fig. 4-1A, B),

HH36 embryos were processed to visualize cartilage. Alcian blue staining of hindlimbs revealed that Dorking autopods had a preaxial ectopic digit (Fig. 4-1C, D). The extra digit was typically fused to the base of the hallux, or first digit ray.

The digits of chicken hindlimbs each have a distinct number of phalanges. By counting the phalanges of a digit ray, identity can be inferred. In the chicken hindlimb, the most anterior digit, digit I, has two phalanges while digit II has three phalanges; this pattern continues through digit IV, which has five phalanges. To further characterize the extra digits in Dorking hindlimbs, the phalanges of the extra digit were counted and compared to wild type (Fig. 4-1C,D and Table 4-1).

To address the issue of variable penetrance, 35 Dorking embryos were analyzed.

In Dorkings, 81% (n=70) of hindlimbs had some degree of polydactyly with no defect observed in any of the forelimbs analyzed (n=70). Of the hindlimbs containing an extra digit, 70% (n=70) resembled a digit II (three phalanges were visible). In instances when a digit II did not occur, ectopic digits resembled a digit I (10%; 7/70 limbs), or a “blip”

(1.4%; 1/70 limbs). Finally, of the 70 hindlimbs analyzed, 19% (n=70) showed no polydactyly and resembled wild type limbs (summarized in Table 4-1). In addition to providing data specific to our population, these data confirm previous observations made 80 years-ago regarding the penetrance of Dorking polydactyly (Punnett and

Pease, 1929).

Ectopic Hedgehog Signaling Occurs in the Anterior of Late Stage Dorking Hindlimbs

To determine if ectopic hedgehog signaling was present in Dorking embryos, Shh

expression was examined using whole mount RNA in situ hybridizations. From HH21 to

HH25, normal posterior expression of Shh was observed in all Dorkings (Fig. 4-2A,B

and data not shown). However at HH26, a small region of ectopic Shh was detected in

the anterior of Dorking hindlimbs (Fig. 4-2C,D). In HH26 hindlimbs, 70% (n=10)

contained ectopic Shh expression. No abnormal Shh expression was observed in the

forelimb at any stage examined (data not shown).

To investigate if the Shh present in the anterior of Dorking hindlimbs was

activating the hedgehog signaling pathway, we examined expression of Ptc1 and Bmp2,

targets of hedgehog signaling (Laufer et al., 1994; Marigo et al., 1996b). At HH21,

normal Ptc1 expression was observed (Fig. 4-2E,F). In the Dorking hindlimb at HH25,

one stage prior to visible ectopic Shh expression, ectopic Ptc1 was observed (Fig. 4-2

G, H). Similar to that observed for Ptc1, normal expression of Bmp2 was detected at

HH21 (Fig. 4-2I,J). At HH26, when ectopic Shh was first observed, Bmp2 was ectopically expressed in the anterior of Dorking hindlimbs (Fig. 4-2K,L). Additional targets of the hedgehog pathway, Gli1 and HoxD13, were also ectopically expressed at

HH26 (Fig. 4-3). These data demonstrated that Shh and hedgehog target genes were ectopically expressed in Dorking hindlimb mesenchyme.

Gene Expression in the AER is Expanded in Dorking Hindlimbs

Fgf4 is normally restricted to the posterior region of the AER, however, anterior expansion of Fgf4 has been observed previously in several mutants with preaxial polydactyly (Masuya et al., 1995; Sharpe et al., 1999). Analysis of Dorking hindlimbs by whole mount RNA in situ hybridization at HH21 revealed that Fgf4 was anteriorly expanded in Dorking hindlimbs (Fig. 4-2M,N). By stage HH25, Fgf4 expression ceased in wild type embryos, however, Fgf4 expression persisted in Dorking hindlimbs (Fig. 4-

2O,P).

A second FGF, Fgf8, has also been shown to be expanded in mouse mutants with

polydactyly (Yada et al., 2002). An analysis of wild type and Dorking limb buds demonstrated that Fgf8 and Sp8, an activator of Fgf8 (Bell et al., 2003; Kawakami et al.,

2004), were slightly expanded in the anterior of Dorking hindlimbs (Fig. 4-3). These data

suggest that, in addition to anterior expansion of Fgf4, the overall length of the AER was expanded in Dorkings.

Ectopic Fgf4 Precedes Ectopic Shh in Dorking Hindlimbs

In situ hybridization data suggested that in Dorking hindlimbs ectopic Fgf expression preceded ectopic expression of Shh and hedgehog target genes by 5 HH stages (approximately 24 hours). One limitation of RNA in situ hybridization is that this

technique cannot detect low levels of gene expression. To confirm that ectopic Fgf4 expression in Dorking hindlimbs preceded ectopic Shh, we used RT-PCR, a more

sensitive technique, to detect Fgf4 and Shh expression in Dorking limbs. Dorking and

wild type hindlimb or forelimb buds at HH21 were removed and trisected into anterior,

middle and posterior segments (Fig. 4-4A). The middle portion of the limb was discarded and gene expression in the anterior and posterior segments was analyzed. In all posterior pools analyzed at HH21, both Shh and Fgf4 were present (Fig. 4-4B). In anterior cDNA pools made from either wild type or Dorking forelimbs, neither Shh nor

Fgf4 was detected. In the hindlimb, cDNA from the anterior of wild type limb buds expressed neither Fgf4 nor Shh transcripts. However, in the Dorking hindlimb Fgf4 but not Shh was detected in the anterior limb bud (Fig. 4-4B). These data confirmed our in situ hybridization experiments and indicated that Fgf4 was ectopically expressed in the anterior of Dorking hindlimbs prior to Shh expression.

Extended Expression of Fgf4 in the Dorking Hindlimb is Maintained Independent of Hedgehog Signaling

In Dorking hindlimbs, we have demonstrated that Fgf4 exhibits a temporal shift in expression. Cyclopamine is a steroidal alkaloid, which inhibits hedgehog signaling by blocking Smoothened (Smo) (Chen et al., 2002; Incardona et al., 1998). To determine if extended Fgf4 expression in Dorking hindlimbs required Shh, hedgehog signaling was inhibited in Dorking hindlimbs using cyclopamine. Treatment of Dorking limbs with cyclopamine resulted in a reduction in Ptc1 expression in the hindlimb (5/6 limbs) whereas none of the mock treated limbs (0/8 limbs) showed reduced Ptc1 expression

(Fig. 4-4C-F). These results indicated that our treatment method was capable of blocking hedgehog signaling in Dorking hindlimbs (p=0.002).

Analysis of different limbs treated in an identical manner revealed that Dorking limbs retained ectopic Fgf4 expression both after treatment with cyclopamine (12/15 limbs) and HBC (8/11 limbs; Fig. 4-4G-J). Comparison of these proportions revealed that the differences were not statistically significant (p=0.69). The treatment of wild type limbs with cyclopamine at a stage when Fgf4 was expected to be expressed resulted in a loss of Fgf4 (p=0.011; data not shown). These data indicated that prolonged maintenance of Fgf4 expression in the AER of Dorking hindlimbs did not require hedgehog signaling.

Cell Death is Decreased in the Anterior Necrotic Zone of Dorking Hindlimbs

Loss of programmed cell death has been observed in chickens with polydactyly

(Dvorak and Fallon, 1991; Hinchliffe and Thorogood, 1974). To investigate apoptosis in

Dorking hindlimbs, we used LysoTracker Red, an acidotropic dye that accumulates in the lysosomes of apoptotic cells and phagocytic cells engulfing apoptotic cells (Zucker et al., 1999). Analysis of the anterior necrotic zone (ANZ) of Dorking hindlimbs after staining by LysoTracker Red revealed that Dorking hindlimbs had a reduced domain of cell death (Fig. 4-5A-D).

To confirm the observed decrease in cell death, we performed whole mount RNA in situ hybridizations using a riboprobe against Dkk1. Dkk1 is a Wnt antagonist, which is spatially and functionally associated with programmed cell death in the developing limb (Grotewold and Ruther, 2002). An examination of Dkk1 expression in Dorking hindlimbs revealed that transcripts were down-regulated in the anterior necrotic zone compared to age matched wild type limb buds (Fig. 4-5E-H). The decrease in Dkk1 expression was consistent with a loss of cell death in the ANZ in Dorking hindlimbs.

Inhibition of Ectopic Fgf but not Hedgehog Signaling in the Dorking Hindlimb can Rescue the Reduction of Cell Death Present in the Anterior Necrotic Zone

Increases to the amount of both FGF and SHH in the anterior of a developing limb have been shown to inhibit cell death in the anterior necrotic zone (Montero et al., 2001;

Sanz-Ezquerro and Tickle, 2000). To separate whether ectopic activation of the Fgf or

Shh signaling pathway played a role in changes to cell death in Dorking hindlimbs, each

pathway was independently inhibited followed by an analysis of cell death. After

inhibition of hedgehog signaling with cyclopamine, 10 of 12 limbs analyzed (83%) had a

domain of cell death that was comparable to Dorkings (Fig. 4-6A-D). The number of

limbs with a Dorking-like domain of cell death after cyclopamine treatment was similar to

untreated limbs (see Table 4-2). These data indicate that inhibition of Shh signaling in

Dorking hindlimbs is not capable of restoring a normal domain of cell death.

To determine if FGF signaling was required to block cell death in the anterior necrotic zone of Dorking hindlimbs, SU5402, a small molecule inhibitor that inhibits all

FGF signaling by blocking tyrosine phosphorylation of FGF receptors was used

(Mohammadi et al., 1997). Inhibition of anterior FGF signaling in Dorking hindlimbs resulted in an increase in the domain of cell death in 50% (n=10) of limbs analyzed (Fig.

4-6E-H). In limbs that had rescued cell death, an increase in LysoTracker staining was observed directly over the SU5402 coated bead. Treatment with a DMSO soaked bead resulted in a Dorking expression pattern in 100% (n=7) of limbs analyzed. These data indicated that treatment with SU5402 had a significant effect on cell death in the developing Dorking hindlimb (p=0.019) and suggests that increases in FGF signaling but not hedgehog signaling are capable of inhibiting cell death in the anterior necrotic zone of Dorkings hindlimbs.

Discussion

Polydactyly is a common congenital malformation with multiple potential causes.

Because human embryos are difficult to obtain for studies of the molecular origins of polydactyly, our knowledge of the molecular etiology of polydactyly relies in part on experiments in model systems. The first recorded description of polydactyly in the chicken model system was in Columella’s Latin text “De Rei Rustica” [translated to

English in (Columella, 1941)]. Although they did not bear the name in Roman times, the polydactylous chickens described in Columella’s text are thought to be Dorkings.

Since the Roman era, Reginald Punnett, among others, have been interested in the genetics of Dorking polydactyly (Darwin, 1890; Punnett and Pease, 1929).

However, prior to the present study the developmental basis of the extra digit in

Dorkings has remained uninvestigated. To better understand the potential origins of polydactyly in vertebrates, we used modern genetic and molecular approaches to investigate the signaling pathways responsible for causing polydactyly in Dorking chickens. Our data indicate that the ectopic preaxial digit found in the Dorking hindlimb initiates from ectopic expression of Fgfs in the AER, which occurs independent of

anterior Shh expression.

Multiple Roles for FGFs in the Formation of Supernumerary Digits in Dorkings

Through gene expression analysis, we found several differences in Dorking

hindlimbs when compared to wild type hindlimbs. Most notable were the differences in

Shh and Fgf expression during hindlimb development. Ectopic Shh is known to be

capable of inducing ectopic Fgf4 (Laufer et al., 1994; Niswander et al., 1994). Our data

revealed that Fgf4 expression preceded Shh expression in the Dorking hindlimb. In

addition, Shh is required for the maintenance of Fgf4 (Chiang et al., 2001). In Dorkings,

treatment with cyclopamine revealed that hedgehog signaling was not required for the maintenance of ectopic Fgf4 expression. These data suggest that Fgf4 and possibly additional factors are ectopically expressed independent of Shh in the anterior of the

Dorking AER.

AER-Fgfs have been hypothesized to establish progenitor numbers in the developing limb (Boulet et al., 2004; Sun et al., 2002). One interesting problem with this

hypothesis is that after total AER-Fgf removal, autopodal skeletal elements are lost,

however, cell death is in a location too proximal to affect autopod progenitor number

(Mariani et al., 2008; Sato et al., 2007). Although the mechanism that causes cell death

in a proximal domain is unclear, it is known that changes in Fgf signaling are capable of

affecting autopod progenitors. For example, partial removal of receptors required for

Fgf signaling results in distal cell death, which causes a loss of autopodal skeletal

elements (Verheyden et al., 2005). Clearly, the location of cell death after changes in

Fgf signaling is important for interpretation of the Dorking phenotype. Based on fate

mapping studies, the decrease in cell death in the anterior necrotic zone observed in

Dorkings occur in a region of the limb where autopod progenitor cells are located

(Vargesson et al., 1997). We propose that changes in cell death present in Dorkings

are capable of altering autopod progenitor cell numbers, resulting in reduced cell death

and polydactyly.

In addition to a role for Fgfs in the inhibition of cell death, Fgfs have been shown to

act as a chemoattractant (Li and Muneoka, 1999). It has been suggested that through

migration of new cells towards the AER, ectopic Fgf expression may result in preaxial

polydactyly (Tanaka et al., 2000). It is possible that in Dorkings, ectopic Fgf expression

attracts cells to the anterior of the developing limb. In combination with a reduction in cell death, attraction of additional cells could greatly increase the number of autopod progenitors in the anterior of the Dorking hindlimb resulting in polydactyly.

In this work, we have demonstrated that inhibiting Fgf signaling in Dorkings results in an increase in cell death. We hypothesize that changes in Fgf signaling in Dorking hindlimbs causes an increase in the number of autopod progenitors. Contrary to this hypothesis, over-expression of AER-Fgfs in mice leads to postaxial polydactyly (Lu et al., 2006), not the preaxial polydactyly found in Dorkings. We propose that these differences may arise because mouse limb buds have a necrotic zone, which appears later and more distally than the anterior necrotic zone in chick limb buds (Fernandez-

Teran et al., 2006). Such differences in cell death between the mouse and chick limb buds may be responsible for the differences in polydactyly observed upon over- expression of Fgfs.

The Role of Shh in Dorking Polydactyly

Our data revealed a role for ectopic Fgfs in initiating Dorking polydactyly. In addition to ectopic Fgf expression, ectopic Shh was also observed. Concentration and time of exposure to a source of SHH protein determines identity of extra digits (Yang et al., 1997). Additionally, the cells that compose a normal digit II in mice have been shown to require response to SHH (Ahn and Joyner, 2004; Harfe et al., 2004). In

Dorkings, variable expression of Shh (see Table 2) offers a potential explanation for the differences observed in digit identity. Because the formation of a normal digit I is independent of SHH (Chiang et al., 1996; Ros et al., 2003) and viral over-expression of

Fgf2 has been shown to result in the formation of extra digit I’s (Riley et al., 1993), we

propose that in embryos, which ectopically express Fgfs but not ectopic Shh, the limb

field is expanded. The increased number of progenitors result in an extra digit I in the absence of Shh. When anterior Shh is expressed in addition to ectopic Fgfs, an extra digit II forms in Dorking hindlimbs.

Ideally, analysis of gene expression in an early limb bud and the resulting skeletal pattern should be analyzed in the same limb. Since it is not feasible to perform gene expression studies on living chick limb buds, we have compared the percentage of embryos containing an ectopic digit II and ectopic Shh expression to examine the role of

Shh in patterning ectopic digits in Dorkings. In both cases, 70% of the analyzed embryos had either an ectopic digit II or ectopic Shh expression in the anterior hindlimb

(see Tables 1 and 2). In Dorkings, 10% of the analyzed skeletal preparations exhibited an ectopic digit I instead of a digit II. In our analysis of Fgf4 expression, we observed

16% more embryos with ectopic Fgf4 expression than embryos with ectopic Shh expression.

While consistent with our hypothesis that formation of an ectopic digit I forms in the absence of ectopic Shh in Dorkings, these data do not rule out the possibility that low levels of Shh may be responsible for polydactyly in Dorking hindlimbs. Additionally,

high doses of transient Shh may be present in embryos where an ectopic digit I was

observed. However, inspection of 56 limbs revealed no Shh expression between HH

stages 20-25. Because we observed changes in cell death prior to detection of Shh or

Ptc1, we favor a model where extra digits in Dorkings are initiated independent of

anterior Shh expression. In this model, ectopic Shh only plays a role in specifying digit II

and not in initiating polydactyly.

Potential Genetic Causes for Dorking Polydactyly

The genetic lesion(s) responsible for formation of the ectopic postaxial digit in

Dorkings is unknown. Our data suggests that ectopic digit formation in Dorkings is due to misexpression of Fgfs in the AER, in particular anterior expansion of Fgf4 within the

AER. The DNA elements responsible for Fgf4 expression have been identified in mouse

(Fraidenraich et al., 1998). By searching the recently completed G. gallus genome, we identified a DNA region homologous to the mouse Fgf4 enhancer. Within this DNA element, ten single nucleotide polymorphisms were identified through comparison of the

Dorking sequence with wild type sequence derived from Red Jungle Fowl (C.B. and

B.D.H., unpublished results). We are currently investigating the potential role of these

SNPs in altering transcription factor binding to the Fgf4 enhancer.

While it is possible that a mutation in the Dorking version of this DNA element is responsible for ectopic Fgf4 expression our data favor a model in which a mutation in a factor upstream of Fgf expression produces the ectopic postaxial digit found in

Dorkings. In part, our hypothesis is based on our findings that both Fgf8 and Sp8 in addition to Fgf4 are ectopically expressed in the AER of Dorking hindlimb buds. Since ectopic Fgf4 expression does not induce ectopic Fgf8 expression (Lu et al., 2006), we propose that a mutation upstream of Fgf expression is most likely responsible for the presence of an ectopic postaxial digit in Dorkings.

Experimental Procedures

Embryo Collection

Dorkings and Dominique chickens used for this study were maintained according to common standards and practices at Morningside Nature Center, Gainesville Florida.

Dominique chickens have a normal number of digits and were used as wild type

controls for gene expression and cell death analysis. Photographs of live roosters were taken on-site with a Nikon Coolpix 7900 digital camera. Eggs were collected every morning by the staff at Morningside Nature Center and stored at room temperature for no longer than a week. In the lab, eggs were incubated at 37°C with supplemental water for humidity. Prior to harvest, eggs were fenestrated with blunted forceps and staged according to the staging guide published by Hamburger and Hamilton (HH)

(Hamburger and Hamilton, 1951).

Skeletal Preparation

For skeletal preparation, HH stage 36 embryos were collected and fixed overnight in 4% formaldehyde in PBS. After evisceration, embryos were incubated in an alcian blue staining solution (70% Ethanol, 30% Acetic Acid and 0.02% w/v Alcian Blue) for 1-

2 days. Embryos were then cleared overnight in 0.5% KOH before being stored in 80% glycerol. After embryos were cleared, phalanges were counted to determine digit identity.

Whole Mount RNA in situ Hybridization

Harvested embryos were incubated overnight in 4% formaldehyde at 4°C. In situ hybridization was performed as described previously (Wilkinson and Neito, 1993).

Plasmids for generating the following riboprobes have been previously described: Shh

(Roelink et al., 1994), Ptc1 (Marigo et al., 1996b), Bmp2 (Francis et al., 1994), Fgf4

(Niswander et al., 1994), Gli1 (Marigo et al., 1996a), HoxD13 (Izpisua-Belmonte et al.,

1991) and Fgf8 (Crossley et al., 1996). Plasmid for generating Sp8 riboprobes was constructed by RT-PCR using the following primers: F 5’-ACCTGCAACAAGATCGGC and R 5’-TCCATGAGGTGCTGCCC.

RNA Isolation, cDNA Synthesis and RT-PCR

Two independent pools of Dorking tissue and one pool of wild type tissue each

consisting of limbs from 10 embryos were collected. Briefly, trisected limbs were

harvested into ice-cold TRIzol reagent (Invitrogen, Carlsbad, CA). RNA was isolated

according to the manufacturer’s protocol. cDNA was synthesized from 2 μg of total

RNA using AMV reverse transcriptase (Roche Applied Science, Indianapolis, IN) and

oligo dT oligonucleotides. Equal quantities of cDNA were used in each RT-PCR

reaction. The primers used for RT-PCR were as follows: Shh (F 5’-

CGGGAGCTGACAGACTGATG, R 5’-CTGATTTCGCTGCCACTGAG), Fgf4 (F 5’-

GCGACTACCTCCTGGGCATC, R 5’-CCGTTTTTGCTCAGGGCTATG) and GAPDH (F

5’-GACGGCAGGTATTAGTTCAAGG, R 5’-ACCATCAAGTCCACAACACGGTTGCTG).

LysoTracker and Drug Treatments

For LysoTracker (Molecular Probes L-7528) analysis, embryos were harvested

into PBS and incubated with 5 µl/ml LysoTracker solution in PBS at 37°C for 30 min.

After staining, embryos were rinsed in PBS, fixed in 4% formaldehyhde in PBS

overnight at 4°C and placed into 100% methanol at –20°C for long-term storage.

Cyclopamine (Sigma Alderich) was dissolved in 45% HBC as described previously

(Incardona et al., 1998). Cyclopamine was administered by separating the amnion adjacent to the hindlimbs using forceps. Sterile pipettes were used to direct 5 µl of HBC or HBC plus cyclopamine (1mg/mL) onto the hindlimbs of HH23 Dorking embryos.

Embryos were then allowed to develop for 20 hours at 37°C and harvested. SU5402

(Merck Chemicals Ltd., Nottingham) was dissolved in DMSO. Ag 1-X2 (Bio-Rad

Laboratories, Hercules, CA) formate-derived beads were incubated in SU5402 (10mM) or DMSO (control) alone for 1hr. Beads were then implanted in the anterior of HH24

Dorking hindlimbs for 10 hours at 37°C. The proportion of drug treated limbs similar to control treated limbs was compared using a z-score. A priori statistical significance was set at a level of 0.05, and all tests were two-sided.

Figure 4-1. Dorking chicken hindlimbs have an ectopic preaxial digit II. Digital images were taken of A) wild type and B) Dorking hindlimbs. In A and B, Digits are numbered according to location. An asterisk denotes the presence of an ectopic digit; “Sp” denotes the location of a spur (keratinous outgrowth), which is present in the hindlimbs of the Dominique roosters (wild type; see Experimental Procedures). Insets in A and B are of the adult wild type Dominique and Dorking chicken, respectively. Skeletal preparations performed on C) wild type and D) Dorking HH36 hindlimbs. Limbs are oriented with the most anterior digit at the top of the panel. In C and D, digit numbers have been assigned according to phalanx number. The ectopic Dorking digit present in panel D resembles a digit II since it has 3 phalanges and is fused to the hallux.

Figure 4-2. Ectopic hedgehog signaling and Fgf expression occurs in the Dorking hindlimb. Whole mount RNA in situ hybridizations using riboprobes against Shh in A) HH21 wild type limbs B) HH21 Dorking hindlimbs, C) HH26 wild type hindlimbs and D) HH26 Dorking hindlimbs. Whole mount RNA in situ hybridizations using riboprobes against Ptc1 in E) HH21 wild type limbs F) HH21 Dorking hindlimbs, G) HH25 wild type hindlimbs and H) HH25 Dorking hindlimbs. Whole mount RNA in situ hybridizations using riboprobes against Bmp2 in I) HH21 wild type limbs J) HH21 Dorking hindlimbs, K) HH25 wild type hindlimbs and L) HH25 Dorking hindlimbs. Whole mount RNA in situ hybridizations using riboprobes against Fgf4 in M) HH21 wild type limbs N) HH21 Dorking hindlimbs, O) HH25 wild type hindlimbs and P) Dorking At HH25. Arrows indicate ectopic expression. Insets in O & P are end on views of hindlimbs.

Figure 4-3. Additional ectopic gene expression in Dorkings. Whole mount RNA in situ hybridizations using riboprobes against Gli1 in A) HH21 wild type limbs B) HH21 Dorking hindlimbs, C) HH26 wild type hindlimbs and D) HH26 Dorking hindlimbs. Whole mount RNA in situ hybridizations using riboprobes against Ptc1 in E) HH21 wild type limbs F) HH21 Dorking hindlimbs, G) HH25 wild HoxD13 hindlimbs and H) HH25 Dorking hindlimbs. Whole mount RNA in situ hybridizations using riboprobes against Fgf8 at HH21 ventral views of I) wild type hindlimbs J) Dorking hindlimbs, and oblique views of K) HH21 wild type hindlimbs and L) HH21 Dorking hindlimbs. Whole mount RNA in situ hybridizations using riboprobes against Sp8 at HH21 ventral views of I) wild type hindlimbs J) Dorking hindlimbs, and oblique views of K) HH21 wild type hindlimbs and L) HH21 Dorking hindlimbs Arrows indicate ectopic expression. Insets in O & P are end on views of hindlimbs. Boxes in K and L and O and P are the same size and were used to demonstrate expansion of Fgf8 and Sp8 in Dorking hindlimbs. The top of the boxes was placed where the anterior limb bud was attached to the body wall, or flank. For all genes examined, no ectopic expression was observed in Dorking forelimbs.

Figure 4-4. Ectopic Fgf4 expression in the Dorking hindlimb precedes and is independent of ectopic Shh. A) Schematic diagram of the strategy used to isolate RNA from forelimbs and hindlimbs. Limbs were trisected and RNA pooled from the anterior (A) and posterior (P) regions of the limb bud (see Experimental Procedures). The middle portion was discarded. B) PCR analysis of Fgf4 and Shh expression in the anterior and posterior of Dorking and wild type fore- and hindlimbs demonstrated that Fgf4 was expressed in the anterior region of the Dorking hindlimb prior to Shh expression (noted by an asterisk). GAPDH was used as a loading control. In situ hybridization analyzing Ptc1 expression in C) wild type hindlimbs, D) untreated Dorking hindlimbs, E) mock treated with 45% HBC Dorking hindlimbs and F) Cyclopamine treated Dorking hindlimbs. In situ hybridization analyzing Fgf4 expression in C) wild type hindlimbs, D) untreated Dorking hindlimbs, E) mock treated with 45% HBC Dorking hindlimbs and F) Cyclopamine treated Dorking hindlimbs. The arrows in D & E marks ectopic expression. Arrows in H-J highlight expression of Fgf4.

Figure 4-5. Cell death is decreased in the anterior necrotic zone (ANZ) of Dorking hindlimbs. Ventral views of LysoTracker red staining in A) wild type and B) Dorking hindlimbs. Anterior views of LysoTracker red staining in C) wild type and D) Dorking hindlimbs. Ventral views of in situ hybridizations against Dkk1 in E) wild type and F) Dorking hindlimbs. Anterior views of in situ hybridizations against Dkk1 expression in G) wild type and H) Dorking hindlimbs.

Figure 4-6. Inhibition of FGF but not Hedgehog signaling rescues cell death in the anterior necrotic zone. Cyclopamine treatment of Dorking hindlimbs followed by LysoTracker Red to visualize cell death. A) Wild type limbs had increased cell death in the anterior necrotic zone compared to either B) untreated or C) mock HBC treated Dorking hindlimbs. D) Treatment of Dorkings hindlimbs with Cyclopamine dissolved in HBC resulted in an inability to rescue cell death in 83% (n=12) of limbs analyzed. To illustrate the domain of cell death in the anterior necrotic zone, white boxes with identical dimensions were placed on the images. SU5402 inhibition of Tyrosine Kinase receptors followed by staining with LysoTracker Red to assay the amount of cell death present in the ANZ. E) Wild type limbs had increased cell death in the anterior necrotic zone compared to either F) untreated or G) mock PBS treated Dorking hindlimbs. H) After implantation of a SU5402 soaked bead, cell death was greatly increased in 56% (n=9) of limbs analyzed. In G and H the location of the beads used in the experiments are represented by white circles. All panels are views of the anterior edge of the hindlimb with the ventral surface facing left.

Table 4-1: Summary of skeletal phenotypes among 35 Dorking incross progeny. Individuals Number of Number of Number of No extra

observed Digit II’s * Digit I’s † Blebs § digit Both Legs with Extra Digit 2’s * 23 46 0 0 0 Both Legs with Extra Digit 1’s † 2 0 4 0 0 One Leg with Extra Digit 2 *, One 2 2 2 0 0 Leg Extra Digit 1 † One Leg with Extra Digit 2 *, One 1 1 0 0 1 Leg with no Extra Digit One Leg with Extra Digit 1 †, One 1 0 1 0 1 Leg with no Extra Digit One Leg with Extra Bleb §, One 1 0 0 1 1 Leg with no Extra Digit No Extra Digits on Either Leg 5 0 0 0 10 Total Number of Limbs 49 7 1 13 Percent of the Total 70% 10% 1.4% 18.6% * A digit II was defined as a supernumerary digit with three cartilaginous condensations. † A digit I was defined as a supernumerary digit with two cartilaginous condensations. § A bleb was defined as a singular supernumerary condensation of cartilage.

Table 4-2: Summary of Variability of Gene Expression, Vital Dye Staining Total Number of Percent of Limbs Marker Analyzed Stage Analyzed Limbs Analyzed Different from Normal Shh HH20-25 56 0.00% HH26 10 70.00%* Ptc1 HH21+ 10 0.00% HH25 8 50.00% Bmp2 HH26 8 25.00% Fgf4 HH21 20 35.00% HH25+ 12 83.33%† LysoTracker HH25 12 83.33%† Dkk1 HH24+ 10 80.00%† * Similar to the percent of limbs with a digit II (70% [49 of 70 limbs]; Table 1). † Similar to the percent of limbs with polydactyly (81.4% [57 of 70 limbs]; Table 1).

CHAPTER 5 CONCLUDING REMARKS

The development of the vertebrate limb requires precise control of cellular and molecular events. Signaling centers required within the developing limb for proper pattern formation have been identified by tissue grafting and removal. More recently, molecules such as Shh and Fgf have been shown to be required for the function of the

ZPA and AER respectively. The field of limb development currently focuses on an integration of the theoretical framework with discoveries elucidating the role of signaling pathways in pattern formation (Chapter 1).

Since its identification by John Saunders and Mary Gasseling, the ZPA has been a focus of research investigating pattern formation in the limb. In this work, we identified a unique area of the limb bud ectoderm that responds to a signal produced from the ZPA

(Chapter 3). We present evidence that the area of hedgehog signaling pathway activation in the ectoderm is required for proper autopod patterning. In addition, we presented evidence that activation of the hedgehog signaling pathway in the AER of the developing vertebrate limb is involved in refining levels of Shh expression from the underlying mesoderm.

In addition to a functional role for hedgehog signaling within the AER, we identified a response to the hedgehog signaling pathway outside of the AER in the ectoderm between the AER and the flank (known as the posterior margin), and in the dorsal and ventral posterior ectoderm. For a more complete understanding of the role hedgehog signaling plays in the ectoderm of the developing limb, we are interested in removing active hedgehog signaling from the remaining portions of the ectoderm. However, no cre recombinase alleles were available at the time of our study to identify a role for

hedgehog signaling in the ectoderm outside the AER. Two available cre alleles, Msx2-

Cre and Brn4-Cre are expressed in the AER and the ventral ectoderm (Ahn et al., 2001;

Sun et al., 2000). We selected Msx2-Cre for analysis in Chapter 3, but these cre alleles maintain gene expression in the dorsal ectoderm and the posterior margin, and are not suitable for a complete ectoderm removal of hedgehog signaling. We also considered the epithelial K14-Cre (Dassule et al., 2000) for use to remove Smo from the entire

ectoderm. However, removal of hedgehog signaling using K14-Cre has been reported

to have no effect on the limb (Gritli-Linde et al., 2007). This could be a result of the limited number of limb cells that express K14-Cre at E10.5 when Shh is expressed (MGI

Database). The RAR-Cre is expressed throughout the limb bud ectoderm.

Unfortunately, this allele is also expressed in the mesoderm of the limb (Barrow et al.,

2003), confounding interpretation of removal of hedgehog signaling from the ectoderm.

A large-scale screening of GLI-binding sites revealed several GLI binding sites

upstream of the Shh target Ptch1 (Vokes et al., 2007). One of these GLI-binding sites,

known as Ptch1:Peak7, was capable of driving expression of a reporter throughout the

ectoderm of the developing limb without inducing expression in the mesoderm

(Appendix A). Using this element to drive expression of cre recombinase, we can

express cre recombinase in any portion of the ectoderm that activates the hedgehog

signaling pathway. Removal of Smo using this cre recombinase from the entire region of

the ectoderm that contains active hedgehog signaling will provide information about the

function of hedgehog signaling in the entire posterior ectoderm.

Implantation of a source of retinoic acid, a known inducer of Shh, in the posterior

limb bud decreases the length of the AER and anterior placement increases the length

of the AER (Lee and Tickle, 1985), suggesting differences in the way that the anterior and posterior portions of the limb respond to Shh. In Chapter 3, we identify active hedgehog signaling in the posterior AER which limits the length of the AER. Although outside of the scope of this work, differences in the ability of the AER to respond to hedgehog signaling could explain differences in response in the anterior and posterior limb to active hedgehog signaling. Because Shh can control growth through the modulation of genes, which promote G1-S transition (Roy and Ingham, 2002), a lack of response in the AER could result in an increase in growth without increased hedgehog signaling restricting the AER, as is the case in the posterior (Chapter 3).

A lack of response to hedgehog signaling in the anterior AER could occur through either restricted diffusion of hedgehog protein to the anterior AER or loss of an essential component of hedgehog signal transduction in the anterior AER. An activated form of

Smoothened (Smo) has been isolated from basal cell carcinomas (Xie et al., 1998). In mice, Smo with an activating mutation has been inserted into the ubiquitously expressed Rosa26 locus. A floxed stop cassette blocks expression in the absence of cre recombinase. However, cre recombinase expression allows constitutive activation of hedgehog signaling (Mao et al., 2006). The ability of the anterior AER to activate hedgehog signaling can be tested by forcing active hedgehog signaling throughout the

AER, and then determining the effect on limb pattern and gene expression. Expression of the activated form of Smo throughout the anterior AER would allow us to determine if the anterior AER was capable of responding to hedgehog signaling. The ability of SHH to diffuse to the anterior AER could then be tested by implanting beads into the anterior

of the limb, and analyzing limbs for SHH protein and targets of hedgehog signaling within the ectoderm.

From early stages of limb condensation to outgrowth from the body wall, many molecules and pathways have been described within the context of limb development.

An important question is how the molecular information within the limb is converted into digit condensations. In culture, limb ectoderm inhibits chondrogenesis (Solursh et al.,

1981). This effect is mediated by WNT and FGF diffusion from the ectoderm, which inhibit the expression of Sox9 (ten Berge et al., 2008). Because Sox9 is required for the

condensation of chondrogenic cells (Barna and Niswander, 2007), cells’ proximity to the

ectoderm, which acts as a source of Wnts and Fgfs, inhibits their ability to form

cartilage.

These findings have interesting implications for the studies reported in this work. In

Chapters 3 and 4, extra condensations form as a result of an increase in AER-Fgfs and

changes in cell death. One effect of a limb field expanded by an increase in proliferation

or a decrease in death is an increase in the volume of cells in the limb. If there is a

threshold distance from the ectoderm for cells to be outside the influence of WNTs and

FGFs to activate Sox9 expression, an increase in limb volume could increase the

number of cells that are an appropriate distance from the developing ectoderm to form

cartilage. After exhaustion of WNTs and FGFs, the increased number of progenitors

could result in the ectopic cartilage condensations observed in Chapters 3 and 4.

In this work, we have identified unique molecular interactions involved in

patterning the vertebrate limb. Activation of hedgehog signaling in the ectoderm

(Chapter 3) and inhibition of cell death in the anterior necrotic zone of Dorking

polydactylous mutants (Chapter 4) represent novel mechanisms for the autopod to refine gene expression, cell number within the limb field, and establish an appropriate array of digits.

APPENDIX A CRE RECOMBINASE EXPRESSION DRIVEN BY THE PEAK7 ELEMENT OF THE PTCH1 PROMOTER

For years, developmental biologists were limited in their study of genes with embryonic lethal phenotypes. To overcome lethality associated with important developmental genes, conditional knockout technology was developed to remove genes from specific tissues (Lewandoski, 2001). Using the bacteriophage protein cre recombinase, sequences of DNA flanked by LoxP sites can be removed in any tissue that expresses the cre recombinase. In general, promoter fragments expressed in tissues of interest are used to drive the expression of cre recombinase (Lewandoski,

2001). A limiting factor for conditional knockout technology is the availability of alleles driving expression of cre recombinase.

As we report in Chapter 3, Shh activates the hedgehog signaling pathway within the posterior ectoderm of the developing vertebrate limb. A functional role for the hedgehog signaling pathway was demonstrated in the AER by removing an essential component of the hedgehog signaling pathway, Smo. Loss of hedgehog signaling from the AER resulted in skeletal defects. Our analysis showed hedgehog signaling pathway activation throughout the posterior ectoderm. Using an AER specific cre recombinase to remove hedgehog signaling from cells in the limb allowed the limb ectoderm, aside from the AER, to respond to hedgehog signaling. A complete analysis of the function of the hedgehog signaling pathway in the developing limb ectoderm requires more complete removal of Smo from the entire ectoderm. Currently, there are no alleles capable of driving expression of cre recombinase in the early ectoderm that are not also expressed in the mesoderm of the developing limb.

Analysis of binding sites of the GLI proteins identified six GLI-binding regions upstream of the transcriptional start of Ptch1 (Vokes et al., 2007). Expanded analysis of

the data set revealed an additional GLI-binding domain. Because it was the seventh

peak of GLI-binding activity this 788bp fragment was named Ptch1:Peak7. Ptch1:Peak7

is located 154.2 Kb upstream of the promoter of Ptch1. Expression of a reporter driven

by the Ptch1:Peak7 regulatory element revealed activation of Ptch1:Peak7 specifically

in the ectoderm at E10.5 (Figure A-1).

The use of an eGFP-cre recombinase fused to an estrogen receptor ligand

binding domain (eEGFPCreErt2) allows temporal control of cre activity by requiring

tamoxifen induced nuclear translocation of the cre recombinase for gene removal

(Lewandoski, 2001). To drive expression of cre recombinase specifically in the

ectoderm of the developing limb, we used the Ptch1:Peak7 element. The eGFPCreErt2

fusion protein has been previously shown to have activity both in vitro and in vivo

(Kobayashi et al., 2008).

Linear constructs were injected by pronuclear injection. After injection into the male pronuclei of embryos, potential founders were identified by PCR genotyping. In total, four founders were identified as positive for the Ptch1:Peak7CreERt2construct.

These founders were crossed to the R26 reporter (R26R) and expression of cre recombinase was confirmed by expression of β-galactosidease. (Soriano, 1999).

Currently, one founder has been analyzed in detail. Embryos produced from this founder with a female positive for the R26R allele treated with tamoxifen (2mg/40g body weight) at E9.5, exhibited β-galactosidase activity in several portions of the embryo at

E11.5 including the lung, the floor plate of the neural tube, a subset of cranial nerves

and the posterior ectoderm of the limb bud (Figure A-2). In combination with the conditional Smo allele, the Ptch1:Peak7CreERt2 allele will allow us to analyze limbs that are unable to respond to hedgehog signaling throughout the posterior ectoderm during most of the developmental stages of the limb. In addition, these transgenics represents the first use of a portion of Ptch1 to activate cre recombinase expression. The

Ptch1:Peak7CreERt2 line can be used in tandem with the Shhgfpcre line (Harfe, 2004)

to analyze gene removal in cells that express Shh and a subset of cells that respond to

hedgehog signaling.

Figure A-1. β-Galactosidase activity driven by the Ptch1:LacZ allele and the Ptch1:Peak7_LacZ allele at E10.5. Comparison of β-Galactosidase in A) whole embryos with Ptch1:LacZ to B) whole embryos with the Ptch1:Peak7_LacZ allele reveal that the Peak7 enhancer element is in a subset of cells which express Ptch1 (detected by β-Galactosidase activity). β- Galactosidase in limb buds is restricted to the posterior of both C) Ptch1:LacZ and D) Ptch1:Peak7_LacZ limbs. Stained sections reveal β-Galactosidase throughout the posterior of E) Ptch1:LacZ limbs but only in the posterior ectoderm of F) Ptch1:Peak7_LacZ limbs. Images provided courtesy of Matthew McFarlane and Andy McMahon.

Figure A-2. Activation of cre recombinase in tissues with both the Ptch1:Peak7CreERt2 and the R26R allele at E11.5. Comparison of whole embryos A) without cre recombinase and B) with Ptch1:Peak7CreERt2reveal cre recombinase expression within the same tissues that express Ptch1:Peak7_LacZ (compare Fig. A-3B to Fig. A-1B). C) Dissection of the embryo revealed cre recombinase activity in the lung and surrounding the neural tube (marked by arrows). D) Dissection of a more caudal portion of the embryo revealed cre recombinase activity in the floor plate of the neural tube and posterior to the neural tube. Cre recombinase activity could also be detected in E) the posterior of the forelimb bud and in a few cells in F) the posterior of the hindlimb bud.

APPENDIX B OLIGONUCLEOTIDES USED AS GENOTYPING PRIMERS

Table B-1. Oligonucleotides used as genotyping primers. Allele Primer Names Tm Mutant Normal Primer Sequence (5’->3’) (°C) Band Band Size Size (bp) (bp) Rosa26R SorGF1 62 250 500 AAAGTCGCTCTGAGTTGTTAT SorGR1 GGAGCGGGAGAAATGGATATG SorGR2 GCGAAGAGTTTGTCCTCAACC Ptch1:LacZ PtchlacZASF1 67 310 GCAAAGACCTCGGGACTCAC PtchlacZASR1 AAGGCGATTAAGTTGGGTAACG Msx2Cre MSX2F2 56 ~500 GACTTTTCAGTTTGGGCG CreB1-MSX AACATCTTCAGGTTCTGCGG Smoflox SmoMutF2 65 ~500 GGAGCTCGAGGAGGGACGTG SmoMutR2 GCGCTACCGGTGGATGTGG Smo IMR1834 62 200 CCACTGCGAGCCTTTGCGCTAC IMR1835 CCCATCACCTCCGCGTCGCA Shhflox Shh1015 65 520 450 ATGCTGGCTCGCCTGGCTGTGGAA Shh1016 GAAGAGATCAAGGCAAGCTCTGGC Shh ShhGenoF 470 GACCCAACTCCGATGTGTTCCGT ShhGenoR2 TCTTCCCTTCATATCTGCCGC Cre BHCreF2 65 260 TGACGGTGGGAGAATGTTAAT BHCreR1 GCCGTAAATCAATCGATGAGT GFP GFP2s 62 570 ATGGTGAGCAAGGGCGAGGAG GFP2aS CGATGGGGGTGTTCTGCTGGTAGT LacZ GeoF 62 270 GTCGTTTTACAACGTCGTGACT GeoR GATGGGCGCATCGTAACCGTGC Ert2Cre ERT2Cre F1 65 400 ATGGATTTCCGTCTCTGGTG ERT2Cre R1 AAGGTTGGCAGCTCTCATGT Ptch1:Peak7_Ert2 Peak7F1 62 500 GCCTCTAATCACCCACCCTTC Peak7R1 GGCTGCTCAGTTTGGATGTTC Ptch1:Peak7_eGFPCre Peak7eGFPF1 62 480 CTGAAGGGCTGGGACTATGA Peak7eGFPR1 AAGTCGTGCTGCTTCATGTG

APPENDIX C PROBES USED FOR RNA IN SITU HYBRIDIZATION

Table C-1. Probes used for RNA in situ hybridization. Gene Name Organism Plasmid Antisense Antisense Number Enzyme Polymerase Smo Mouse BH185 SalI T7 Ptch1 Mouse BH141 BamHI T3 Sox9 Mouse BH165 HindIII T7 Fgf8 Mouse BH99 XhoI Sp6 Fgf8 Chicken BH12 EcoRI T7 Fgf4 Chicken BH11 EcoRI T7 Shh Mouse BH39 HindIII T3 Fgf4 Mouse BH144 BamHI T3 Gremlin1 Mouse BH75 PstI T3 Shh Chicken BH16 SalI Sp6 Ptc1 Chicken BH17 SalI T3 Bmp2 Chicken BH18 HindIII T3 Gli1 Chicken BH100 SacI T7 HoxD13 Chicken BH167 EcoRV T3 Sp8 Chicken BH30 NotI T3 Dkk1 Chicken BH151 NcoI Sp6

LIST OF REFERENCES

Ahn, K., Mishina, Y., Hanks, M.C., Behringer, R.R., and Crenshaw, E.B., 3rd (2001). BMPR-IA signaling is required for the formation of the apical ectodermal ridge and dorsal-ventral patterning of the limb. Development 128, 4449-4461.

Ahn, S., and Joyner, A.L. (2004). Dynamic changes in the response of cells to positive hedgehog signaling during mouse limb patterning. Cell 118, 505-516.

Akiyama, H., Chaboissier, M.C., Martin, J.F., Schedl, A., and de Crombrugghe, B. (2002). The transcription factor Sox9 has essential roles in successive steps of the chondrocyte differentiation pathway and is required for expression of Sox5 and Sox6. Genes Dev 16, 2813-2828.

Altabef, M., Clarke, J.D., and Tickle, C. (1997). Dorso-ventral ectodermal compartments and origin of apical ectodermal ridge in developing chick limb. Development 124, 4547- 4556.

Barna, M., and Niswander, L. (2007). Visualization of cartilage formation: insight into cellular properties of skeletal progenitors and chondrodysplasia syndromes. Dev Cell 12, 931-941.

Barrow, J.R., Thomas, K.R., Boussadia-Zahui, O., Moore, R., Kemler, R., Capecchi, M.R., and McMahon, A.P. (2003). Ectodermal Wnt3/beta-catenin signaling is required for the establishment and maintenance of the apical ectodermal ridge. Genes Dev 17, 394-409.

Bell, S.M., Schreiner, C.M., Goetz, J.A., Robbins, D.J., Scott, W.J., Jr., Bell, S.M., Schreiner, C.M., and Scott, W.J. (2005). Shh signaling in limb bud ectoderm: potential role in teratogen-induced postaxial ectrodactyly. Dev Dyn 233, 313-325.

Bell, S.M., Schreiner, C.M., and Scott, W.J. (1999). Disrupting the establishment of polarizing activity by teratogen exposure. Mech Dev 88, 147-157.

Bell, S.M., Schreiner, C.M., Waclaw, R.R., Campbell, K., Potter, S.S., and Scott, W.J. (2003). Sp8 is crucial for limb outgrowth and neuropore closure. Proc Natl Acad Sci U S A 100, 12195-12200.

Benazet, J.D., Bischofberger, M., Tiecke, E., Goncalves, A., Martin, J.F., Zuniga, A., Naef, F., and Zeller, R. (2009). A self-regulatory system of interlinked signaling feedback loops controls mouse limb patterning. Science 323, 1050-1053.

Biesecker, L.G. (2002). Polydactyly: how many disorders and how many genes? Am J Med Genet 112, 279-283.

Bitgood, M.J., and McMahon, A.P. (1995). Hedgehog and Bmp genes are coexpressed at many diverse sites of cell-cell interaction in the mouse embryo. Dev Biol 172, 126- 138.

Bouldin, C.M., and Harfe, B.D. (2009). Aberrant FGF signaling, independent of ectopic hedgehog signaling, initiates preaxial polydactyly in Dorking chickens. Dev Biol 334, 133-141.

Boulet, A.M., Moon, A.M., Arenkiel, B.R., and Capecchi, M.R. (2004). The roles of Fgf4 and Fgf8 in limb bud initiation and outgrowth. Dev Biol 273, 361-372.

Chan, D.C., Laufer, E., Tabin, C., and Leder, P. (1995). Polydactylous limbs in Strong's Luxoid mice result from ectopic polarizing activity. Development 121, 1971-1978.

Charite, J., de Graaff, W., Shen, S., and Deschamps, J. (1994). Ectopic expression of Hoxb-8 causes duplication of the ZPA in the forelimb and homeotic transformation of axial structures. Cell 78, 589-601.

Charite, J., McFadden, D.G., and Olson, E.N. (2000). The bHLH transcription factor dHAND controls Sonic hedgehog expression and establishment of the zone of polarizing activity during limb development. Development 127, 2461-2470.

Chen, J.K., Taipale, J., Cooper, M.K., and Beachy, P.A. (2002). Inhibition of Hedgehog signaling by direct binding of cyclopamine to Smoothened. Genes Dev 16, 2743-2748.

Chiang, C., Litingtung, Y., Harris, M.P., Simandl, B.K., Li, Y., Beachy, P.A., and Fallon, J.F. (2001). Manifestation of the limb prepattern: limb development in the absence of sonic hedgehog function. Dev Biol 236, 421-435.

Chiang, C., Litingtung, Y., Lee, E., Young, K.E., Corden, J.L., Westphal, H., and Beachy, P.A. (1996). Cyclopia and defective axial patterning in mice lacking Sonic hedgehog gene function. Nature 383, 407-413.

Cohn, M.J. (2000). Developmental biology. Giving limbs a hand. Nature 406, 953-954.

Columella, L.J.M. (1941). On agriculture, with a recension of the text and an English translation, Vol II (Cambridge, Massachusetts, Harvard University Press).

Crossley, P.H., and Martin, G.R. (1995). The mouse Fgf8 gene encodes a family of polypeptides and is expressed in regions that direct outgrowth and patterning in the developing embryo. Development 121, 439-451.

Crossley, P.H., Minowada, G., MacArthur, C.A., and Martin, G.R. (1996). Roles for FGF8 in the induction, initiation, and maintenance of chick limb development. Cell 84, 127-136.

Dahn, R.D., Davis, M.C., Pappano, W.N., and Shubin, N.H. (2007). Sonic hedgehog function in chondrichthyan fins and the evolution of appendage patterning. Nature 445, 311-314.

Darwin, C. (1900). The variation of animals and plants under domestication. New York: D. Appleton and Co.

Dassule, H.R., Lewis, P., Bei, M., Maas, R., and McMahon, A.P. (2000). Sonic hedgehog regulates growth and morphogenesis of the tooth. Development 127, 4775- 4785.

Dudley, A.T., Ros, M.A., and Tabin, C.J. (2002). A re-examination of proximodistal patterning during vertebrate limb development. Nature 418, 539-544.

Dvorak, L., and Fallon, J.F. (1991). Talpid2 mutant chick limb has anteroposterior polarity and altered patterns of programmed cell death. Anat Rec 231, 251-260.

Fallon, J.F., Lopez, A., Ros, M.A., Savage, M.P., Olwin, B.B., and Simandl, B.K. (1994). FGF-2: apical ectodermal ridge growth signal for chick limb development. Science 264, 104-107.

Fernandez-Teran, M., Piedra, M.E., Kathiriya, I.S., Srivastava, D., Rodriguez-Rey, J.C., and Ros, M.A. (2000). Role of dHAND in the anterior-posterior polarization of the limb bud: implications for the Sonic hedgehog pathway. Development 127, 2133-2142.

Fernandez-Teran, M.A., Hinchliffe, J.R., and Ros, M.A. (2006). Birth and death of cells in limb development: a mapping study. Dev Dyn 235, 2521-2537.

Fraidenraich, D., Lang, R., and Basilico, C. (1998). Distinct regulatory elements govern Fgf4 gene expression in the mouse blastocyst, myotomes, and developing limb. Dev Biol 204, 197-209.

Francis, P.H., Richardson, M.K., Brickell, P.M., and Tickle, C. (1994). Bone morphogenetic proteins and a signalling pathway that controls patterning in the developing chick limb. Development 120, 209-218.

Fromental-Ramain, C., Warot, X., Messadecq, N., LeMeur, M., Dolle, P., and Chambon, P. (1996). Hoxa-13 and Hoxd-13 play a crucial role in the patterning of the limb autopod. Development 122, 2997-3011.

Fuse, N., Maiti, T., Wang, B., Porter, J.A., Hall, T.M., Leahy, D.J., and Beachy, P.A. (1999). Sonic hedgehog protein signals not as a hydrolytic enzyme but as an apparent ligand for patched. Proc Natl Acad Sci U S A 96, 10992-10999.

Galloway, J.L., Delgado, I., Ros, M.A., and Tabin, C.J. (2009). A reevaluation of X- irradiation-induced phocomelia and proximodistal limb patterning. Nature 460, 400-404.

Goetz, J.A., Suber, L.M., Zeng, X., and Robbins, D.J. (2002). Sonic Hedgehog as a mediator of long-range signaling. Bioessays 24, 157-165.

Goodrich, L.V., Milenkovic, L., Higgins, K.M., and Scott, M.P. (1997). Altered neural cell fates and medulloblastoma in mouse patched mutants. Science 277, 1109-1113.

Gritli-Linde, A., Hallberg, K., Harfe, B.D., Reyahi, A., Kannius-Janson, M., Nilsson, J., Cobourne, M.T., Sharpe, P.T., McMahon, A.P., and Linde, A. (2007). Abnormal hair development and apparent follicular transformation to mammary gland in the absence of hedgehog signaling. Dev Cell 12, 99-112.

Gritli-Linde, A., Lewis, P., McMahon, A.P., and Linde, A. (2001). The whereabouts of a morphogen: direct evidence for short- and graded long-range activity of hedgehog signaling peptides. Dev Biol 236, 364-386.

Grotewold, L., and Ruther, U. (2002). The Wnt antagonist Dickkopf-1 is regulated by Bmp signaling and c-Jun and modulates programmed cell death. Embo J 21, 966-975.

Hamburger, V., and Hamilton, H.L. (1951). A Series Of Normal Stages In The Development Of The Chick Embryo. Journal Of Morphology 88, 49-&.

Haramis, A.G., Brown, J.M., and Zeller, R. (1995). The limb deformity mutation disrupts the SHH/FGF-4 feedback loop and regulation of 5' HoxD genes during limb pattern formation. Development 121, 4237-4245.

Harfe, B.D., Scherz, P.J., Nissim, S., Tian, H., McMahon, A.P., and Tabin, C.J. (2004). Evidence for an expansion-based temporal Shh gradient in specifying vertebrate digit identities. Cell 118, 517-528.

Harrison, R.G. (1918). Experiments on the development of the fore limb of Amblystoma, a self-differentiating equipotential system. Journal of Experimental Zoology 25, 413-461.

Helms, J.A., Kim, C.H., Eichele, G., and Thaller, C. (1996). Retinoic acid signaling is required during early chick limb development. Development 122, 1385-1394.

Hinchliffe, J.R., and Thorogood, P.V. (1974). Genetic inhibition of mesenchymal cell death and the development of form and skeletal pattern in the limbs of talpid3 (ta3) mutant chick embryos. J Embryol Exp Morphol 31, 747-760.

Hooper, J.E., and Scott, M.P. (2005). Communicating with Hedgehogs. Nat Rev Mol Cell Biol 6, 306-317.

Incardona, J.P., Gaffield, W., Kapur, R.P., and Roelink, H. (1998). The teratogenic Veratrum alkaloid cyclopamine inhibits sonic hedgehog signal transduction. Development 125, 3553-3562.

Izpisua-Belmonte, J.C., Tickle, C., Dolle, P., Wolpert, L., and Duboule, D. (1991). Expression of the homeobox Hox-4 genes and the specification of position in chick wing development. Nature 350, 585-589.

Kawakami, Y., Capdevila, J., Buscher, D., Itoh, T., Rodriguez Esteban, C., and Izpisua Belmonte, J.C. (2001). WNT signals control FGF-dependent limb initiation and AER induction in the chick embryo. Cell 104, 891-900.

Kawakami, Y., Esteban, C.R., Matsui, T., Rodriguez-Leon, J., Kato, S., and Belmonte, J.C. (2004). Sp8 and Sp9, two closely related buttonhead-like transcription factors, regulate Fgf8 expression and limb outgrowth in vertebrate embryos. Development 131, 4763-4774.

Kengaku, M., Capdevila, J., Rodriguez-Esteban, C., De La Pena, J., Johnson, R.L., Izpisua Belmonte, J.C., and Tabin, C.J. (1998). Distinct WNT pathways regulating AER formation and dorsoventral polarity in the chick limb bud. Science 280, 1274-1277.

Khokha, M.K., Hsu, D., Brunet, L.J., Dionne, M.S., and Harland, R.M. (2003). Gremlin is the BMP antagonist required for maintenance of Shh and Fgf signals during limb patterning. Nat Genet 34, 303-307.

Kobayashi, A., Valerius, M.T., Mugford, J.W., Carroll, T.J., Self, M., Oliver, G., and McMahon, A.P. (2008). Six2 defines and regulates a multipotent self-renewing nephron progenitor population throughout mammalian kidney development. Cell Stem Cell 3, 169-181.

Kocsis, E., Trus, B.L., Steer, C.J., Bisher, M.E., and Steven, A.C. (1991). Image averaging of flexible fibrous macromolecules: the clathrin triskelion has an elastic proximal segment. J Struct Biol 107, 6-14.

Laufer, E., Nelson, C.E., Johnson, R.L., Morgan, B.A., and Tabin, C. (1994). Sonic hedgehog and Fgf-4 act through a signaling cascade and feedback loop to integrate growth and patterning of the developing limb bud. Cell 79, 993-1003.

Lee, J., and Tickle, C. (1985). Retinoic acid and pattern formation in the developing chick wing: SEM and quantitative studies of early effects on the apical ectodermal ridge and bud outgrowth. J Embryol Exp Morphol 90, 139-169.

Lettice, L.A., Heaney, S.J., Purdie, L.A., Li, L., de Beer, P., Oostra, B.A., Goode, D., Elgar, G., Hill, R.E., and de Graaff, E. (2003). A long-range Shh enhancer regulates expression in the developing limb and fin and is associated with preaxial polydactyly. Hum Mol Genet 12, 1725-1735.

Lettice, L.A., Horikoshi, T., Heaney, S.J., van Baren, M.J., van der Linde, H.C., Breedveld, G.J., Joosse, M., Akarsu, N., Oostra, B.A., Endo, N., et al. (2002). Disruption of a long-range cis-acting regulator for Shh causes preaxial polydactyly. Proc Natl Acad Sci U S A 99, 7548-7553.

Lewandoski, M. (2001). Conditional control of gene expression in the mouse. Nat Rev Genet 2, 743-755.

Lewandoski, M., Sun, X., and Martin, G.R. (2000). Fgf8 signalling from the AER is essential for normal limb development. Nat Genet 26, 460-463.

Lewis, P.M., Dunn, M.P., McMahon, J.A., Logan, M., Martin, J.F., St-Jacques, B., and McMahon, A.P. (2001). Cholesterol modification of sonic hedgehog is required for long- range signaling activity and effective modulation of signaling by Ptc1. Cell 105, 599-612.

Li, S., and Muneoka, K. (1999). Cell migration and chick limb development: chemotactic action of FGF-4 and the AER. Dev Biol 211, 335-347.

Lu, H.C., Revelli, J.P., Goering, L., Thaller, C., and Eichele, G. (1997). Retinoid signaling is required for the establishment of a ZPA and for the expression of Hoxb-8, a mediator of ZPA formation. Development 124, 1643-1651.

Lu, P., Minowada, G., and Martin, G.R. (2006). Increasing Fgf4 expression in the mouse limb bud causes polysyndactyly and rescues the skeletal defects that result from loss of Fgf8 function. Development 133, 33-42.

Maatouk, D.M., Choi, K.S., Bouldin, C.M., and Harfe, B.D. (2009). In the limb AER Bmp2 and Bmp4 are required for dorsal-ventral patterning and interdigital cell death but not limb outgrowth. Dev Biol 327, 516-523.

MacCabe, J.A., Saunders, J.W., Jr., and Pickett, M. (1973). The control of the anteroposterior and dorsoventral axes in embryonic chick limbs constructed of dissociated and reaggregated limb-bud mesoderm. Dev Biol 31, 323-335.

Mackem, S., and Lewandoski, M. (2009). Limb development takes a measured step toward systems analysis. Sci Signal 2, pe33.

Maden, M. (1982). Vitamin A and pattern formation in the regenerating limb. Nature 295, 672-675.

Mahmood, R., Bresnick, J., Hornbruch, A., Mahony, C., Morton, N., Colquhoun, K., Martin, P., Lumsden, A., Dickson, C., and Mason, I. (1995). A role for FGF-8 in the initiation and maintenance of vertebrate limb bud outgrowth. Curr Biol 5, 797-806.

Mao, J., Ligon, K.L., Rakhlin, E.Y., Thayer, S.P., Bronson, R.T., Rowitch, D., and McMahon, A.P. (2006). A novel somatic mouse model to survey tumorigenic potential applied to the Hedgehog pathway. Cancer Res 66, 10171-10178.

Mao, J., McGlinn, E., Huang, P., Tabin, C.J., and McMahon, A.P. (2009). Fgf-dependent Etv4/5 activity is required for posterior restriction of Sonic Hedgehog and promoting outgrowth of the vertebrate limb. Dev Cell 16, 600-606.

Mariani, F.V., Ahn, C.P., and Martin, G.R. (2008). Genetic evidence that FGFs have an instructive role in limb proximal-distal patterning. Nature 453, 401-405.

Marigo, V., Johnson, R.L., Vortkamp, A., and Tabin, C.J. (1996a). Sonic hedgehog differentially regulates expression of GLI and GLI3 during limb development. Dev Biol 180, 273-283.

Marigo, V., Scott, M.P., Johnson, R.L., Goodrich, L.V., and Tabin, C.J. (1996b). Conservation in hedgehog signaling: induction of a chicken patched homolog by Sonic hedgehog in the developing limb. Development 122, 1225-1233.

Martin, G.R. (1998). The roles of FGFs in the early development of vertebrate limbs. Genes Dev 12, 1571-1586.

Masuya, H., Sagai, T., Moriwaki, K., and Shiroishi, T. (1997). Multigenic control of the localization of the zone of polarizing activity in limb morphogenesis in the mouse. Dev Biol 182, 42-51.

Masuya, H., Sagai, T., Wakana, S., Moriwaki, K., and Shiroishi, T. (1995). A duplicated zone of polarizing activity in polydactylous mouse mutants. Genes Dev 9, 1645-1653.

Mercader, N., Leonardo, E., Azpiazu, N., Serrano, A., Morata, G., Martinez, C., and Torres, M. (1999). Conserved regulation of proximodistal limb axis development by Meis1/Hth. Nature 402, 425-429.

Mercader, N., Leonardo, E., Piedra, M.E., Martinez, A.C., Ros, M.A., and Torres, M. (2000). Opposing RA and FGF signals control proximodistal vertebrate limb development through regulation of Meis genes. Development 127, 3961-3970.

Mohammadi, M., McMahon, G., Sun, L., Tang, C., Hirth, P., Yeh, B.K., Hubbard, S.R., and Schlessinger, J. (1997). Structures of the tyrosine kinase domain of fibroblast growth factor receptor in complex with inhibitors. Science 276, 955-960.

Montero, J.A., Ganan, Y., Macias, D., Rodriguez-Leon, J., Sanz-Ezquerro, J.J., Merino, R., Chimal-Monroy, J., Nieto, M.A., and Hurle, J.M. (2001). Role of FGFs in the control of programmed cell death during limb development. Development 128, 2075-2084.

Murone, M., Rosenthal, A., and de Sauvage, F.J. (1999). Sonic hedgehog signaling by the patched-smoothened receptor complex. Curr Biol 9, 76-84.

Nissim, S., Hasso, S.M., Fallon, J.F., and Tabin, C.J. (2006). Regulation of Gremlin expression in the posterior limb bud. Dev Biol 299, 12-21.

Niswander, L. (2002). Interplay between the molecular signals that control vertebrate limb development. Int J Dev Biol 46, 877-881.

Niswander, L., Jeffrey, S., Martin, G.R., and Tickle, C. (1994). A positive feedback loop coordinates growth and patterning in the vertebrate limb. Nature 371, 609-612.

Niswander, L., Tickle, C., Vogel, A., Booth, I., and Martin, G.R. (1993). FGF-4 replaces the apical ectodermal ridge and directs outgrowth and patterning of the limb. Cell 75, 579-587.

Ohuchi, H., Nakagawa, T., Yamamoto, A., Araga, A., Ohata, T., Ishimaru, Y., Yoshioka, H., Kuwana, T., Nohno, T., Yamasaki, M., et al. (1997). The mesenchymal factor, FGF10, initiates and maintains the outgrowth of the chick limb bud through interaction with FGF8, an apical ectodermal factor. Development 124, 2235-2244.

Panman, L., Galli, A., Lagarde, N., Michos, O., Soete, G., Zuniga, A., and Zeller, R. (2006). Differential regulation of gene expression in the digit forming area of the mouse limb bud by SHH and gremlin 1/FGF-mediated epithelial-mesenchymal signalling. Development 133, 3419-3428.

Pajni-Underwood, S., Wilson, C.P., Elder, C., Mishina, Y., and Lewandoski, M. (2007). BMP signals control limb bud interdigital programmed cell death by regulating FGF signaling. Development. 134. 2359-2368.

Pearse, R.V., 2nd, Vogan, K.J., and Tabin, C.J. (2001). Ptc1 and Ptc2 transcripts provide distinct readouts of Hedgehog signaling activity during chick embryogenesis. Dev Biol 239, 15-29.

Punnett, R.C., and Pease, M.S. (1929). Genetic studies in poultry VII Notes in polydactyly. Journal Of Genetics 21, 341-366.

Quirk, J., van den Heuvel, M., Henrique, D., Marigo, V., Jones, T.A., Tabin, C., and Ingham, P.W. (1997). The smoothened gene and hedgehog signal transduction in Drosophila and vertebrate development. Cold Spring Harb Symp Quant Biol 62, 217- 226.

Rasband, W.S., ImageJ, U. S. National Institue of Health, Bethesda, Maryland, USA, http://rsb.info.nih.gov/ij/ (1997-2007).

Riddle, R.D., Johnson, R.L., Laufer, E., and Tabin, C. (1993). Sonic hedgehog mediates the polarizing activity of the ZPA. Cell 75, 1401-1416.

Riley, B.B., Savage, M.P., Simandl, B.K., Olwin, B.B., and Fallon, J.F. (1993). Retroviral expression of FGF-2 (bFGF) affects patterning in chick limb bud. Development 118, 95- 104.

Robert, B. (2007). Bone morphogenetic protein signaling in limb outgrowth and patterning. Dev Growth Differ 49, 455-468.

Roelink, H., Augsburger, A., Heemskerk, J., Korzh, V., Norlin, S., Ruiz i Altaba, A., Tanabe, Y., Placzek, M., Edlund, T., Jessell, T.M., et al. (1994). Floor plate and motor neuron induction by vhh-1, a vertebrate homolog of hedgehog expressed by the notochord. Cell 76, 761-775.

Ros, M.A., Dahn, R.D., Fernandez-Teran, M., Rashka, K., Caruccio, N.C., Hasso, S.M., Bitgood, J.J., Lancman, J.J., and Fallon, J.F. (2003). The chick oligozeugodactyly (ozd) mutant lacks sonic hedgehog function in the limb. Development 130, 527-537.

Rowe, D.A., Cairns, J.M., and Fallon, J.F. (1982). Spatial and temporal patterns of cell death in limb bud mesoderm after apical ectodermal ridge removal. Dev Biol 93, 83-91.

Roy, S., and Ingham, P.W. (2002). Hedgehogs tryst with the cell cycle. J Cell Sci 115, 4393-4397.

Sanz-Ezquerro, J.J., and Tickle, C. (2000). Autoregulation of Shh expression and Shh induction of cell death suggest a mechanism for modulating polarising activity during chick limb development. Development 127, 4811-4823.

Sato, K., Koizumi, Y., Takahashi, M., Kuroiwa, A., and Tamura, K. (2007). Specification of cell fate along the proximal-distal axis in the developing chick limb bud. Development 134, 1397-1406.

Saunders, J.W. (1948). The Proximo-Distal Sequence Of Origin Of The Parts Of The Chick Wing And The Role Of The Ectoderm. Journal Of Experimental Zoology 108, 363- 403.

Saunders, J.W., and Gasseling, M.T. (1968). Ectodermal-Mesenchymal Interactions in the Origin of Limb Symmetry. In Epithelial-Mesenchymal Interactions; 18th Hahnemann Symposium, R. Fleischmajer, and R.E. Billingham, eds. (Baltimore, Williams & Wilkins), pp. 78-97.

Scherz, P.J., Harfe, B.D., McMahon, A.P., and Tabin, C.J. (2004). The limb bud Shh-Fgf feedback loop is terminated by expansion of former ZPA cells. Science 305, 396-399.

Sesgin, M.Z., and Stark, R.B. (1961). The incidence of congenital defects. Plast Reconstr Surg Transplant Bull 27, 261-267.

Sharpe, J., Lettice, L., Hecksher-Sorensen, J., Fox, M., Hill, R., and Krumlauf, R. (1999). Identification of sonic hedgehog as a candidate gene responsible for the polydactylous mouse mutant Sasquatch. Curr Biol 9, 97-100.

Solursh, M., Singley, C.T., and Reiter, R.S. (1981). The influence of epithelia on cartilage and loose connective tissue formation by limb mesenchyme cultures. Dev Biol 86, 471-482.

Soriano, P. (1999). Generalized lacZ expression with the ROSA26 Cre reporter strain. Nat Genet 21, 70-71.

St-Jacques, B., Hammerschmidt, M., and McMahon, A.P. (1999). Indian hedgehog signaling regulates proliferation and differentiation of chondrocytes and is essential for bone formation. Genes Dev 13, 2072-2086.

Stratford, T., Horton, C., and Maden, M. (1996). Retinoic acid is required for the initiation of outgrowth in the chick limb bud. Curr Biol 6, 1124-1133.

Summerbell, D., Lewis, J.H., and Wolpert, L. (1973). Positional information in chick limb morphogenesis. Nature 244, 492-496.

Sun, X., Lewandoski, M., Meyers, E.N., Liu, Y.H., Maxson, R.E., Jr., and Martin, G.R. (2000). Conditional inactivation of Fgf4 reveals complexity of signalling during limb bud development. Nat Genet 25, 83-86.

Sun, X., Mariani, F.V., and Martin, G.R. (2002). Functions of FGF signalling from the apical ectodermal ridge in limb development. Nature 418, 501-508.

Tabin, C., and Wolpert, L. (2007). Rethinking the proximodistal axis of the vertebrate limb in the molecular era. Genes Dev 21, 1433-1442.

Tanaka, M., Cohn, M.J., Ashby, P., Davey, M., Martin, P., and Tickle, C. (2000). Distribution of polarizing activity and potential for limb formation in mouse and chick embryos and possible relationships to polydactyly. Development 127, 4011-4021. ten Berge, D., Brugmann, S.A., Helms, J.A., and Nusse, R. (2008). Wnt and FGF signals interact to coordinate growth with cell fate specification during limb development. Development 135, 3247-3257.

Tickle, C. (1981). The number of polarizing region cells required to specify additional digits in the developing chick wing. Nature 289, 295-298.

100

Todt, W.L., and Fallon, J.F. (1984). Development of the apical ectodermal ridge in the chick wing bud. J Embryol Exp Morphol 80, 21-41.

Towers, M., Mahood, R., Yin, Y., and Tickle, C. (2008). Integration of growth and specification in chick wing digit-patterning. Nature 452, 882-886. van den Akker, E., Fromental-Ramain, C., de Graaff, W., Le Mouellic, H., Brulet, P., Chambon, P., and Deschamps, J. (2001). Axial skeletal patterning in mice lacking all paralogous group 8 Hox genes. Development 128, 1911-1921.

Vargesson, N., Clarke, J.D., Vincent, K., Coles, C., Wolpert, L., and Tickle, C. (1997). Cell fate in the chick limb bud and relationship to gene expression. Development 124, 1909-1918.

Verheyden, J.M., Lewandoski, M., Deng, C., Harfe, B.D., and Sun, X. (2005). Conditional inactivation of Fgfr1 in mouse defines its role in limb bud establishment, outgrowth and digit patterning. Development 132, 4235-4245.

Verheyden, J.M., and Sun, X. (2008). An Fgf/Gremlin inhibitory feedback loop triggers termination of limb bud outgrowth. Nature 25, 25.

Vokes, S.A., Ji, H., McCuine, S., Tenzen, T., Giles, S., Zhong, S., Longabaugh, W.J., Davidson, E.H., Wong, W.H., and McMahon, A.P. (2007). Genomic characterization of Gli-activator targets in sonic hedgehog-mediated neural patterning. Development 134, 1977-1989.

Vokes, S.A., Ji, H., Wong, W.H., and McMahon, A.P. (2008). A genome-scale analysis of the cis-regulatory circuitry underlying sonic hedgehog-mediated patterning of the mammalian limb. Genes Dev 22, 2651-2663.

Wellik, D.M., Hawkes, P.J., and Capecchi, M.R. (2002). Hox11 paralogous genes are essential for metanephric kidney induction. Genes Dev 16, 1423-1432.

Wilkinson, D.G., and Nieto, M.A. (1993). Detection of messenger RNA by in situ hybridization to tissue sections and whole mounts. Methods Enzymol 225, 361-373.

Wolpert, L. (1969). Positional information and the spatial pattern of cellular differentiation. J Theor Biol 25, 1-47.

Wolpert, L., Tickle, C., and Sampford, M. (1979). The effect of cell killing by x-irradiation on pattern formation in the chick limb. J Embryol Exp Morphol 50, 175-193.

Xie, J., Murone, M., Luoh, S.M., Ryan, A., Gu, Q., Zhang, C., Bonifas, J.M., Lam, C.W., Hynes, M., Goddard, A., et al. (1998). Activating Smoothened mutations in sporadic basal-cell carcinoma. Nature 391, 90-92.

101

Yada, Y., Makino, S., Chigusa-Ishiwa, S., and Shiroishi, T. (2002). The mouse polydactylous mutation, luxate (lx), causes anterior shift of the anteroposterior border in the developing hindlimb bud. Int J Dev Biol 46, 975-982.

Yang, Y., Drossopoulou, G., Chuang, P.T., Duprez, D., Marti, E., Bumcrot, D., Vargesson, N., Clarke, J., Niswander, L., McMahon, A., et al. (1997). Relationship between dose, distance and time in Sonic Hedgehog-mediated regulation of anteroposterior polarity in the chick limb. Development 124, 4393-4404.

Yang, Y., and Niswander, L. (1995). Interaction between the signaling molecules WNT7a and SHH during vertebrate limb development: dorsal signals regulate anteroposterior patterning. Cell 80, 939-947.

Yokouchi, Y., Sasaki, H., and Kuroiwa, A. (1991). Homeobox gene expression correlated with the bifurcation process of limb cartilage development. Nature 353, 443- 445.

Zguricas, J., Heus, H., Morales-Peralta, E., Breedveld, G., Kuyt, B., Mumcu, E.F., Bakker, W., Akarsu, N., Kay, S.P., Hovius, S.E., et al. (1999). Clinical and genetic studies on 12 preaxial polydactyly families and refinement of the localisation of the gene responsible to a 1.9 cM region on chromosome 7q36. J Med Genet 36, 32-40.

Zhang, X.M., Ramalho-Santos, M., and McMahon, A.P. (2001). Smoothened mutants reveal redundant roles for Shh and Ihh signaling including regulation of L/R asymmetry by the mouse node. Cell 105, 781-792.

Zhang, Z., Verheyden, J.M., Hassell, J.A., and Sun, X. (2009). FGF-regulated Etv genes are essential for repressing Shh expression in mouse limb buds. Dev Cell 16, 607-613.

Zhu, J., Nakamura, E., Nguyen, M.T., Bao, X., Akiyama, H., and Mackem, S. (2008). Uncoupling Sonic hedgehog control of pattern and expansion of the developing limb bud. Dev Cell 14, 624-632.

Zucker, R.M., Hunter, E.S., 3rd, and Rogers, J.M. (1999). Apoptosis and morphology in mouse embryos by confocal laser scanning microscopy. Methods 18, 473-480.

Zuniga, A., Michos, O., Spitz, F., Haramis, A.P., Panman, L., Galli, A., Vintersten, K., Klasen, C., Mansfield, W., Kuc, S., et al. (2004). Mouse limb deformity mutations disrupt a global control region within the large regulatory landscape required for Gremlin expression. Genes Dev 18, 1553-1564.

Zwilling, E. (1964). Development of Fragmented and of Dissociated Limb Bud Mesoderm. Dev Biol 89, 20-37.

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K14-Cre recombinase expression in E10.5 limbs was retrieved from the Mouse Genome Database (MGD), (K14-Cre MGI ID:2445832), Mouse Genome Informatics, The Jackson Laboratory, Bar Harbor, Maine. World Wide Web (URL: http://www.informatics.jax.org). [January, 2010]

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BIOGRAPHICAL SKETCH

Cortney Bouldin earned his Bachelor of Science in microbiology and cell sciences from the University of Florida in 2003. Prior to graduate school, he worked on the function of F1F0 ATP synthase in E. coli and the effects of protein restricted diets on the methylation status of DNA within developing embryos. During his graduate career,

Cortney studied pattern formation in the developing vertebrate limb. Specifically, he identified a role for Shh signaling in the ectoderm of the developing limb and studied the molecular mechanisms involved in the formation of polydactyly. In his research, Cortney applied a methodology developed by Eva Kocsis to measure the lengths of protein molecules in electron micrographs to measure the lengths of Apical Ectodermal Ridges in the limb bud. Additionally, he participated in the creation of a cre recombinase allele, which is expressed in a subset of cells that express the hedgehog target Ptch1. Cortney has been an active member in the Society for Developmental Biology where he has received travel awards to attend annual meetings and awards in best student poster competitions. He also attended the 10th International Congress on Limb Development and Regeneration in Madrid, Spain where he gave an oral presentation. Cortney recieved his Ph.D. from the University of Florida in the Spring of 2010, and intends to continue his work investigating mechanisms of pattern formation with a special focus on the early steps of embryo development including gastrulation and axis elongation.

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