ROLE OF ROBO GENES DURING MURINE VERTEBRAL COLUMN FORMATION
By
LISA Y. LAWSON
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
2017
2
© 2017 Lisa Y. Lawson
3
To my parents for their encouragement and support, and to my sister who kept me sane in all my endeavors
4
ACKNOWLEDGMENTS
I thank members of the Cohn Lab for their support and understanding, and for
being so inclusive in their science and friendship. I thank Kendra McKee for always
being so present, even when we couldn‟t be on the same floor. I couldn‟t have asked for
a better friend and more helpful lab mom. I also thank Emily Merton for being so
generous in her understanding while I worked to finish my studies in the Harfe lab.
Finally, I thank Brian Harfe for his patience and understanding every time I found myself
flustered by a deadline.
5
TABLE OF CONTENTS
page AWKNOWLEDGEMENTS………………………………………………………… 4
LIST OF TABLES……………………………………………………….…………. 7
LIST OF FIGURES……………………………………………………….…….….. 8
ABSTRACT……………………………………………………………………….… 10
CHAPTER
1 LITERATURE REVIEW: DEVELOPMENTAL MECHANISMS OF VERTEBRAL COLUMN AND INTERVERTEBRAL DISC MORPHOGENESIS………………………………………………………
12
Introduction………………………………………………………………... 12 Embryonic Origins of the Vertebral Column…………………….…….. 14 The Notochord………………………………………………….. 15 Somites………………………………………………………….. 16 Developmental Origins of the Annulus Fibrosus……………………… 18 Genes Required For Development of the Nucleus Pulposus……….. 19 Hedgehog Signaling Pathway…………………………………. 19 Foxa1 and Foxa2……………………………………………….. 21 T-Brachyury……………………………………………………… 22 Noto……………………………….………………………………. 22 Genes Required For Development of the Annulus Fibrosus……….. 23 Pax1 and Pax9………………………………………………….. 24 Bapx1……………………………….……………………………. 26 Sox Family of Genes…………………………………………... 26 Scleraxis And TGFβ Signaling………………………………… 28 Conclusions……………………………….………………………………. 29 Genes Required For Murine Vertebral Column Formation…………... 31 Anatomy Of The Vertebral Column And Intervertebral Disc…………. 34 Vertebral Column Morphogenesis In Mice…………………………….. 34 Postnatal Intervertebral Disc Development……………………………. 35 2 ROLE OF ROBO GENES DURING MURINE VERTEBRAL
COLUMN MORPHOGENESIS…………………………………………. 36
Introduction……………………………….……………………………… 36 Results……………………………….…………………………………… 39 Complementary Robo and Slit Expression Patterns in the
Vertebral Column……………………………………………….. 39
Robo expression…………………………………………. 39 Slit expression……………………………………………. 41
6
Intervertebral Disc Malformations………………………………. 42 The Notochord to Nucleus Pulposus Transition...……………. 43 Rib and Sternal Development…………………………………... 48 Proliferative Deficits in the Sclerotome………………………… 49 Robo and Pax1 Expression in the Vertebral Column………… 50 Robo Gene Expression and Function in the Growth Plates…. 52 Annulus Fibrosus Cell Morphology and Gene Expression…... 54 Cartilage Matrix Protein Expression …………………………… 54 Tenascin C Upregulation………………………………………… 55 Discussion and Conclusions…………………………………………….. 56 Role of Robo1 and Robo2 in IVD Development……………… 56 Robo1 and Robo2 Function in the Sclerotome……………….. 59 Robo1 and Robo2 in Growth Plate Maintenance…………….. 60 Figures……………………………………………………………………... 62 3 WNT/β-CATENIN SIGNALING DURING INTERVERTEBRAL DISC
DEVELOPMENT………………………………………………………….. 79
Introduction…………...…………………………………………………… 79 Results……………………………………………………………………... 81 Conclusions……………………………………………………………….. 83 Figures……………………………………………………………………... 86 4 METHODS……………………………….………………………………...
90
Histology……………………………….………………………………….. 90 Skeletal Preparations…………………………………………………….. 90 Western Blot Analysis……………………………………………………. 91 ECM Analysis By Immunofluorescence………………………………... 92 Characterization Of Robo And Slit Expression………………………... 93 X-Gal Staining…………………………………………………… 93 RNA In Situ Hybridization………………………………………. 94 Lineage Tracing Analyses……………………………………………….. 96 Analysis Of Sclerotomal Proliferation…………………………………... 97 EdU Pulse And Detection………………………………………. 97 B-Gal Antibody As Proxy For Robo Expression……………... 98 Quantification……………………………….…………………… 98 Growth Plate Analysis……………………………….…………………… 99 Characterization of Wnt/β-Catenin Signaling Activity………………… 99 Immunofluorescence And Western Blot Antibodies…………………... 100 Genotyping Primers……………………………….……………………… 100
REFERENCES……………………………….…………………………………….. 102 BIOGRAPHICAL SKETCH………………………………………………………...
114
7
LIST OF TABLES
Table Page
1-1 Genes Required For Murine Vertebral Column Formation……… 31
4-1 ECM Immunofluorescence And Western Blot Antibodies……….. 100
4-2 Genotyping Primers…………………………………………………. 100
8
LIST OF FIGURES
Figure page
1-1 Anatomy of the Vertebral Column and Intervertebral Disc…….... 34
1-2 Vertebral Column Development In Mice…………………………... 34
1-3 Postnatal Intervertebral Disc Development……………………….. 35
2-1 Robo1 and Robo2 Expression Analysis by RNA In Situ Hybridization and Xgal Staining…………………………………….
62
2-2 Robo Gene Expression During Vertebral Column Formation....... 63
2-3 Slit Gene Expression During Vertebral Column Formation……... 64
2-4 Removal of Robo1 and Robo2 Causes Intervertebral Disc Malformations in P0 Mice……………………………………………
65
2-5 Conditional Inactivation of Robo2F on Robo1 Null Background.. 66
2-6 The Notochord to Nucleus Pulposus Transition Occurs Normally in the Absence of Robo1 and Robo2………………………………
67
2-7 Lineage Tracing Analysis of ShhCreERT2 Marked Cells............. 68
2-8 Robo1;Robo2 Null Mice Have Deficits in the Distal Ribs and Sterna at P0……………………………………………………….....
69
2-9 Impaired Proliferation in Robo1; Robo2 Null Mutants at 13.5dpc 70
2-10 Robo and Pax1 Have Overlapping Expression Patterns in the Developing Vertebral Column……………………………………….
72
2-11 Disrupted Growth Plate Marker Expression in Robo1;Robo2 Null Mutants…………………...……………………………………...
73
2-12 Robo1;Robo2 Null Annuli Fibrosi Have Aberrant Cell Morphologies And Ectopic Expression of Growth Plate Chondrocyte Markers………………………………………………..
75
2-13 Robo1 and Robo2 Loss is Associated With Reduced Expression of Cartilage Matrix Proteins…………………………...
76
9
2-14 Tenascin C is Upregulated in the Intervertebral Discs of Robo1;Robo2 Null Mice…………………………………………….
77
2-15 Slit1;Slit2 Null Phenotype………...………...………...………....... 78
3-1 WNT/β-Catenin Signaling in the Notochord at 12.5 dpc………… 86
3-2 WNT Signaling in the Embryonic Intervertebral Discs…………… 87
3-3 Postnatal Wnt Signaling in the Intervertebral Discs……………… 88
10
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
ROLE OF ROBO GENES DURING MURINE VERTEBRAL COLUMN FORMATION
By
Lisa Y. Lawson
May 2017
Chair: Brian D. Harfe Major: Medical Sciences – Genetics
The vertebral column is comprised of ossified vertebrae which are adjoined by
the fibrocartilagenous intervertebral discs. In humans, common pathologies affecting the
vertebral column include scoliosis and intervertebral disc degeneration, which is thought
to be a major cause of lower back pain in adults worldwide. The molecular mechanisms
underlying scoliosis and disc degeneration in humans are not well understood. Robo
proteins are a conserved class of transmembrane receptors which are most classically
known for their role in regulating axonal projections via a chemorepulsive mechanism
during nervous system development. The diverse roles of Robo proteins in modulating
cell proliferation, survival, migration, and adhesion in vivo during vertebrate
embryogenesis, however, has been illuminated more clearly by recent studies in Robo
knockout mice. A role for Robo proteins in sclerotome differentiation and vertebral
column development has not been reported previously. Here we show that Robo1 and
Robo2 are expressed in the embryonic sclerotome, and mice that lack both Robo1 and
Robo2 develop vertebral columns with enlarged intervertebral discs with aberrant
annulus fibrosus cell morphology. In addition, we demonstrate that in the absence of
Robo1 and Robo2, the organized expression of Col10a and Ihh in vertebral growth plate
11
chondrocytes is disrupted. These changes are preceded by decreased proliferation in
the sclerotome and are accompanied by deficits in the distal ribs and sterna at birth.
12
CHAPTER 1 LITERATURE REVIEW: DEVELOPMENTAL MECHANISMS OF VERTEBRAL
COLUMN AND INTERVERTEBRAL DISC MORPHOGENESIS
Introduction
The axial skeleton is comprised of the skull and ossified elements of the middle
ear, the hyoid bone at the base of the neck, the ribs, sternum, and the vertebral column
(White, et al. 2012). Within the vertebral column, the intervertebral discs are the
fibrocartilagenous joint-like structures that connect and cushion the vertebrae. In young
and healthy animals, the intervertebral discs are comprised of three functionally distinct
regions that interact synergistically to facilitate fluid and painless movement of the
spine. Each intervertebral disc is comprised of a gel-like core called the nucleus
pulposus, a fibrocartilagenous surrounding called the annulus fibrosus, and the
cartilaginous endplates (Fig. 1-1).
Within each disc, the nucleus pulposus is comprised of cells suspended in a gel-
like matrix of negatively charged proteoglycans that attract and retain water molecules
in the intervertebral disc core (Sivan, et al. 2014). Hydration in the nucleus pulposus
allows for uniform re-distribution of compressive forces generated by vertebral column
movement (Sivan, et al. 2012). Surrounding each nucleus pulposus core is the annulus
fibrosus, which is comprised of spindle-shaped chondrocyte-like cells embedded in a
highly organized matrix of fibrillar proteins. The concentric, lamellar organization and
structured orientation of fibrillar proteins in the annulus fibrosus allow it to withstand the
shear and ply tensions it is subjected to as the intervertebral discs bear loads (Cortes
and Elliot, 2012; Romgens, et al. 2013; Hayes, et al. 2011). Finally, as the intervertebral
discs are aneural and avascular in adulthood, the cartilaginous endplates are important
not only for adhering the discs to adjacent vertebrae but also for their role in nutrient
13
and waste diffusion into and out of the discs (Richardson, et al., 2012; Smith and Elliot,
2011; Malandrino, et al., 2014).
As animals age, cell senescence and metabolic changes in the discs can lead to
disc degeneration and consequently, to impaired disc function and disc related lower
back pain (Colombier, et al., 2014; Gruber et al., 2007; Gruber, et al., 2009; Le Maitre,
et al., 2007; Smith, et al., 2011). Disc degeneration can be caused by metabolic
imbalances or by acute trauma. Hallmarks of disc degeneration include decreased
nucleus pulposus hydration, decreased disc height, ruptured annuli fibrosi, and calcified
endplates (Smith, et al., 2011). A major constituent of the non-collagenous extracellular
matrix in the discs is Aggrecan. Aggrecan plays an integral role in maintaining the load-
bearing properties of the discs, and in humans Aggrecan loss is associated with disc
degenration (Sivan, et al. 2014). By covalently interacting with 100 or more negatively
charged sulfated glycosaminoglycan (GAG) side chain groups, Aggrecan maintains the
osmolality required to keep the discs hydrated (Kiani, et al., 2002).
In humans, Aggrecan abundance in the discs peaks in the early twenties and
steadily declines thereafter (Sivan, et al. 2014). As a consequence of Aggrecan loss,
the ability of nuclei pulposi to uniformly redistribute compressive forces becomes
compromised, leading to uneven pressures against the surrounding annuli fibrosi. As a
result, the annulus fibrosus can tear, leading to eruption of nucleus pulposus matter
through the annulus fibrosus. These tears trigger wound-healing processes that
promote abnormal neo-vascularization and innervation in the discs, which are thought to
be a contributing factor in disc related back pain in humans (Tolofari, et al., 2010;
Freemont, et al., 2002).
14
Disc degeneration and associated back pain in humans is proposed to arise from
a combination of genetic and environmental factors, which can include occupation,
pregnancy, smoking, and body mass index (Abbas, et al., 2013; Dario, et al., 2015;
Hestbaek, et al., 2004; Nasto, et al., 2014). By some estimates, the direct and indirect
economic burden associated with low back pain in the United States alone exceeds
$100 billion annually (Andersson, 1999; Katz, 2006; Buchbinder, et al., 2013). Based on
recent projections made by the Global Burden of Disease Study (GBD), low back pain is
now the leading cause of disability worldwide, superseding heart disease, diabetes, and
major depressive and anxiety disorders (Millenium, 2003). The lifetime prevalence for
low back pain has been reported to be as high as 80-85% (Buchbinder, et al. 2013).
Disc associated lower back pain can be debilitating, limiting range of motion and
quality of life. To reverse disc pathology in the clinic, a better understanding of disc
biology and development in the context of vertebral column formation is necessary. This
literature review focuses on what is known about the molecular and signaling
mechanisms underlying vertebral column formation and intervertebral disc
morphogenesis with an emphasis on genetic mouse models.
Embryonic Origins of the Vertebral Column
The vertebral column forms from cells contained in two distinct embryonic
compartments: the notochord and somites. The notochord gives rise to the nuclei
pulposi of each intervertebral disc in adult mice (Choi and Harfe, 2011; MacCann, et al.,
2012). Within each somite, a specific compartment known as the sclerotome
differentiates to form the vertebrae and annuli fibrosi. Notochord and somite
differentiation occur in unison during embryogenesis. Because signaling interactions
between the notochord and somites are required for sclerotome induction and
15
subsequent development, genetic abnormalities that impair notochord formation impede
vertebral column development early in development.
The Notochord
In mice the notochord forms from the node in the axial mesoderm beginning
around 7.5 dpc (days post coitum) as a tube-like structure that elongates rostro-caudally
at the midline, ventral to the neural tube (Sulik, et al., 1994; Yamanaka, et al., 2007).
The notochord is composed of large vacuolated cells which are enveloped in an
acellular membrane known as the “notochordal sheath” (Paavola, et al. 1980). These
large vacuoles inside notochordal cells are thought to play a role in body axis elongation
by exerting a hydrostatic pressure against the basement membrane of the notochordal
sheath (Corallo, et al., 2015; Stemple, 2005). The acellular notochordal sheath is
composed of collagens, cytokeratins, laminin, fibronectin, and other glycosaminoglycan-
modified proteoglycans (Lehtonen, et al., 1995; Gotz, et al., 1995). The notochordal
sheath, which physically segregates notochordal cells from the surrounding paraxial
mesoderm, is integral to notochord cell survival and is likely synthesized by notochordal
cells. Genetic perturbations that impair notochordal sheath integrity have been linked to
aberrant notochord cell death in mice (Smits and Lefebvre, 2003; Choi and Harfe,
2011). Presently, it is unclear whether cell death occurs as a consequence of
notochordal sheath loss, or vice versa.
Recently, it was proposed that the notochordal sheath was essential for the
notochord to nucleus pulposus transition. In this model, the notochordal sheath acts as
a membranous barrier, separating the axial and paraxial mesoderms (i.e. notochord and
somites, respectively) as notochordal cells move into nucleus pulposus anlagen
16
beginning around 12.5 dpc (Fig. 1-2). In these experiments, Hedgehog signaling was
conditionally removed in the notochord, resulting in disrupted notochordal sheath
integrity, followed by aberrant scattering of notochordal cells throughout the vertebral
column. As a result of Hedghog signaling removal, mice developed to 18.5 dpc with
severe loss of disc and vertebral structures in the axial skeleton (Choi and Harfe, 2011).
Similarly, inactivation of Sox5 and Sox6 together produced defects in the notochordal
sheath followed by deficits in vertebral structures (Smits et al. 2003). Deletion of the
Foxa transcription factors (Foxa1 and Foxa2), which act upstream of Shh, have also
been shown to cause notochordal sheath defects with resultant phenotypes in the
vertebral column (Maier and Harfe, 2013).
Somites
Somites are bilateral blocks of mesoderm that flank the notochord and neural
tube. During embryogenesis cells in the unsegmented pre-somitic mesoderm (PSM)
epithelize and bud off rostrocaudally to form somites, which flank the notochord and
neural tube. (Tam and Trainor, 1994). Somite formation, or somitogenesis, is regulated
by the FGF, Notch, and Wnt signaling pathways (Saga, 2012). In mice, 65 somite pairs
give rise to 65 vertebrae, while in humans 33 somite pairs form the 33 vertebrae
(Theiler, 1972; Haque, et al. 2001).
Once formed, somites acquire dorso-ventral polarity in response to signals from
the surface ectoderm and ventral midline (floor plate and notochord). Wnt signals from
the surface ectoderm induce dermamyotome specification in the dorsal somite. The
dermamyotome, which expresses Pax3, Pax7, and Sim1 (Goulding, et al. 1991; Fan, et
al. 1994), subsequently differentiates to form the myotome and dermatome, which
17
develop into muscles and dermis, respectively. Myotome specification is distinguished
by the expression of Myod1 and Myf5 (Borycki, et al. 1999), which are required to
initiate FGF signaling in the myotome (Brent, et al. 2005). FGF signaling in the myotome
is required for specification of tendon progenitors in the ventral somitic mesoderm
(Brent, et al. 2003).
In the ventral somite, Shh from the notochord and floor plate induce Pax1
expression, which marks specification of the sclerotome. Following the induction of
chondroprogenitors in the sclerotome, cells in the dorsolateral edge of each sclerotome
express Scx (Sclerxis) in response to FGF signaling from the myotome (Brent, et al.
2003). The syndetome is comprised of Scx-expressing cells, which are the progenitors
that develop into tendons and ligaments (Brent, et al. 2003; Brent et al. 2005; Cserjesi,
et al. 1995). Mutations in Scx or its targets, Tenomodulin and Mohawk, produce deficits
in tendon formation or function, including those that anchor the vertebrae to each other
(Shukunami, et al. 2006; Liu, et al. 2015; Murchison, et al. 2007). Syndetome induction
in chick embryos is mediated in part by FREK and FGF effectors Pea3 and Ets (Brent,
et al. 2004).
Finally, the sclerotome gives rise to all the cartilaginous and ossified components
of the ribs and vertebral column. The sclerotome is molecularly distinguishable within
the somitic mesoderm by Pax1, Pax9, and Bapx1 expression (Herbrand, et al. 2002;
Rodrigo, et al. 2003; Peters, et al. 1999). The sclerotome forms the outer parts of the
intervertebral discs, including the annuli fibrosi and cartilaginous endplates (Bruggeman,
et al. 2012). The sclerotome also gives rise to the vertebrae, including the ventral
vertebral bodies the dorsal spinous processes that encircle and protect the spinal cord.
18
The sclerotome also gives rise to the proximal and distal ribs (Aoyama, et al. 2005).
Formation of the vertebrae occurs by re-segmentation, wherein the caudal half of one
somite interacts with the rostral half of an adjacent somite to form one vertebral unit.
Re-segmentation has been observed in chicks by tracking dye-labeled cells and in mice
by observing deficits that occur when genes that regulate somite polarity and re-
segmentation, like Uncx4.1 and Tbx18, are removed (Aoyama, et al. 2000; Leitges, et
al. 2000; Bussen, et al. 2004).
Developmental Origins of the Annulus Fibrosus
The intervertebral discs form from cells present in the notochord and sclerotome
(Choi, et al. 2012; Christ, et al. 2004). All cells in nuclei pulposi are derived from the
notochord (Choi and Harfe, 2011; McCann, et al. 2012). The annulus fibrosi and
endplates form from the sclerotome (Bruggeman, et al. 2012). Cell tracing analyses of
lipophilic dye injections have demonstrated that in chickens, which do not contain nuclei
pulposi, the intervertebral discs (annulus fibrosus) are derived from rostral sclerotome
(Bruggeman, et al. 2012). Other lines of evidence suggesting a sclerotomal origin for
the annuli fibrosi include reports that Pax1 is expressed in the sclerotome and annuli
fibrosi but is absent in the notochord and nuclei pulposi (DiPaola, et al. 2005;
Senthinathan, et al. 2012).
Other genes expressed in the sclerotome include Uncx4.1 (Uncx) and Tbx18
(Leitges, et al. 2000; Mansouri, et al. 1997; Kraus, et al. 2001). Analyses of Uncx4.1-
LacZ transgenic mice showed that Uncx expressing cells in the caudal sclerotome gave
rise to the presumptive endplates and annulus fibrosi of the intervertebral discs
(Takahashi, et al. 2013). Tbx18 was shown to be expressed in the anterior mouse
19
sclerotome at 10.5 dpc and fate mapping of Tbx18Cre;R26R mice showed that these
Tbx18-expressing cells later formed the annulus fibrosus (Kraus, et al. 2001). Both
Uncx and Tbx18 null mice die perinatally with severe abnormalities in the vertebral
column suggestive of defects in lateral sclerotome differentiation (Bussen, et al. 2004;
Leitges, et al. 2000).
Genes Required For Development Of The Nucleus Pulposus
Uncovering the molecular pathways responsible for formation of the
intervertebral discs has lagged compared to many other tissues. For example, the role
Shh plays in limb development was initially described >20 years ago but only recently
was Shh shown to be important for formation of the intervertebral discs (Riddle, et al.
1993; Choi and Harfe, 2011). Many of the key signaling pathways involved in vertebral
column formation and intervertebral disc morphogenesis also play critical roles in
multiple other developmental processes. (Chiang, et al. 1996). Thus, the creation of null
alleles of many of these genes results in early lethality, prior to disc formation.
Advances in mouse genes in the past decade, however, including the creation of large
numbers of conditional mouse alleles and the characterization of Cre alleles, including
ShhCre and NotoCre, have allowed researchers in the field to study the roles of specific
genes in the relevant tissues that give rise to the discs (Harfe, et al. 2004; Choi and
Harfe, 2011; McCann, et al. 2012).
Hedgehog Signaling Pathway
Shh (Sonic Hedgehog) is a secreted ligand that is responsible for activating
Hedgehog signaling within many cell types. It is expressed in the node, notochord and
nucleus pulposus (Choi and Harfe, 2011; Jeong, et al. 2003; Dahia, et al. 2012).
20
Mutations in Shh affect vertebral column formation, both directly and indirectly (Choi
and Harfe, 2011; Chiang, et al. 1996; Choi, et al. 2012). Shh null embryos developed a
node and caudal notochord, as evidenced by the presence of T-brachyury mRNA, but
the notochord quickly disintegrated following down-regulation in Foxa2 in the
rudimentary notochord (Chiang, et al. 1996). These findings suggested that Shh was
dispensable for node formation and initiation of the notochord from the node, but was
required for notochord growth and elongation in a Foxa2-dependent manner (Chiang, et
al. 1996). Shh null embryos were embryonic lethal with severe deficits in multiple organ
systems and had severely reduced axial skeletons with completely absent intervertebral
discs and vertebrae (Chiang, et al. 1996).
Subsequent studies in conditional Hedgehog mutants further illuminated the role
of Shh in notochord and intervertebral disc biology. Hedgehog signaling is mediated by
the Smo (Smoothened) cell surface receptor. Smo removal has been shown to abolish
Hedgehog signaling in targeted cells (Zhang, et al. 2001). When Hedgehog signaling
was conditionally inactivated in the notochord using a floxed allele of Smo, the vertebral
column failed to form (Choi and Harfe, 2011). Hedgehog inactivation in the notochord
resulted in loss of the notochordal sheath, proliferative deficits in the notochord, and
aberrant scattering of notochordal cells throughout the paraxial mesenchyme (Choi and
Harfe, 2011). Hedgehog signaling has also been documented in the postnatal discs and
has been suggested to play a role in re-activating Wnt signaling for intervertebral disc
maintenance in aged animals (Dahia, et al. 2009; Dahia, et al. 2012; Winkler, et al.
2014).
21
Foxa1 and Foxa2
Foxa2 (also called HNF-3b or hepatocyte nuclear factor 3b) is a DNA-binding
protein in the Forkhead box family of transcription factors that is critical for notochord
formation and maintenance (Weinstein, et al. 1994; Ang and Rossant, 1994). Foxa2 null
embryos were described as having defects in primitive streak elongation which resulted
in notochord agenesis followed by embryonic lethality by 11.5 dpc (Ang and Rossant,
1994). Expression of another Foxa family transcription factor, Foxa1 (also called HNF-
3a) has also been described in the notochord (Kaestner, et al. 1994). Homozygous
deletion of Foxa1 did not affect notochord formation and intervertebral disc
morphogenesis (Kaestner, et al. 1994).
A subsequent study illustrated the functional redundancy between Foxa1 and
Foxa2 in the notochord. ShhCre was used to conditionally inactivate a floxed allele of
Foxa2 on a Foxa1 null background in mice (Maier et al. 2013). Using this approach, it
was shown that while removal of Foxa1 or Foxa2 resulted in normal development,
removal of both Foxa1 and Foxa2 in the notochord severely impaired the ability of the
notochord to transition into nuclei pulposi (Maier, et al. 2013). Perinatal Foxa1;Foxa2
mutants had underdeveloped vertebral columns with severely reduced and misshapen
intervertebral discs (Maier, et al. 2013). These results make sense in light of other
studies that showed direct binding of Foxa transcription factors to the regulatory
sequences of many notochord-specific genes (Tamplin, et al. 2011).
Notably, the Foxa1;Foxa2 mutant phenotype was similar to the phenotype
observed in the vertebral column when Hedgehog signaling was inactivated in the
notochord (Maier, et al. 2013; Choi and Harfe, 2011). These results were congruent with
22
what is known about the genetic interaction between Shh and the Foxa transcription
factors. Foxa1 protein has been shown to directly bind the Shh notochord-specific
enhancer (Jeong and Epstein, 2003). Consistent with these observations, Hedgehog
signaling was down-regulated in Foxa1;Foxa2 mutants (Maier, et al. 2013). Based on
these studies, it is likely that the Foxa family of transcription factors directly regulate Shh
expression in the notochord. Curiously, Foxa2 expression was observed in Shh null
embryos. However, in the absence of Shh, Foxa2 was subsequently down-regulated in
the notochord (Chiang, et al. 1996). These results suggest that maintenance of Foxa2
expression is regulated by a Shh-dependent feedback loop in the notochord.
T-Brachyury
T-Brachyury is a conserved T-box transcription factor that is expressed in the
node, notochord, and nucleus pulposus (Wilkinson, et al. 1990; Hermann, 1992; Dahia,
et al. 2012). Compared to Foxa2 null embryos, which failed to form any notochord, T-
brachyury null embryos formed a caudal notochord but died by 10.5 dpc with a
truncated notochord at the forelimb level (Weinstein, et al. 1994; Ang and Rossant,
1994; Beddington, et al. 1992; Rashbass, et al. 1994). During normal development, T-
brachyury is expressed in the notochord and its expression is maintained in notochord-
derived nucleus pulposus cells postnatally (Dahia, et al. 2012). The role of T-brachyury
in postnatal intervertebral discs is unknown. Gene duplications in T-brachyury have
been linked to risk for familial chordomas in humans (Yang, et al. 2009).
Noto
The homeobox transcription factor Noto is expressed in the node and notochord
between 7.5dpc and 12.5 dpc. (Zizic-Mitrecic, et al. 2010; Abdelkhalek, et al. 2004). In
23
mice, Noto is required for tail bud morphogenesis and Noto deletion resulted in
notochord truncation at the tail level (Zizic-Mitrecic, et al. 2010; Abdelkhalek, et al.
2004). Mutations in Noto underlie the floating head (flh) phenotype in zebrafish, which is
characterized by complete absence of a notochord (Talbot, et al. 1995).
In mice Noto is thought to be an important regulator of axial mesoderm
(notochord) identity (McCann, et al. 2012; Zizic-Mitrecic, et al. 2010; Abdelkhalek, et al.
2004; Yamanaka, et al. 2007). In mice where notochord cells had been GFP-labeled,
Noto inactivation resulted in ectopic localization of GFP-labeled cells in the paraxial
mesoderm rather than at the midline, as in control mice. The authors‟ interpretation was
that in the absence of Noto, axial mesoderm (notochord cells) had acquired somitic
mesoderm cell fate and were thus found in the paraxial mesoderm (McCann, et al.
2012). Additional studies, perhaps by probing for Shh, T-brachyury, or Foxa2
expression in these ectopic GFP-positive cells, would enhance our understanding about
the role of Noto in axial mesoderm maintenance. Absence of Shh, T-brachyury, and
Foxa2 in ectopic GFP-positive cells would conclusively demonstrate that a true cell fate
re-specification of notochord cells had occurred in Noto null mutants.
Genes Required For Development Of The Annulus Fibrosis
Following somitogenesis, cells in the ventral somite undergo an epithelial to
mesenchymal transition to form the sclerotome (Hay, 2005; Yusuf and Brand-Saberi,
2006; Christ and Scaal, 2008). In addition to producing the ossified vertebral
components in the axial skeleton, progenitors contained in the embryonic sclerotome
give rise to the cartilaginous annuli fibrosi of each intervertebral disc.
24
Pax1 and Pax9
One of the earliest markers of sclerotome specification is Pax1, which requires
Shh and Noggin expression in the ventral midline for its induction (Wallin, et al. 1994;
McMahon, et al. 1998; Koseki, et al. 1993). Soon after Pax1 induction, Pax9 expression
is activated in the sclerotome (Neubuser, et al. 1995). Pax1 and Pax9 have overlapping
and spatially distinct expression patterns in the sclerotome (Neubuser, et al. 1995).
Following induction of both Pax1 and Pax9, Pax1 expression becomes enriched in the
ventral sclerotome while Pax9 expression becomes relegated to the dorsal sclerotome
(Neubuser, et al. 1995). In the absence of Pax1, the Pax9 expression domain becomes
upregulated and spatially expands in the sclerotome (Peters, et al. 1999).
The role of Pax1 in vertebral column formation is well documented (Table 1-1).
The characterization of multiple Pax1 mutants have illustrated its importance in
sclerotome differentiation. In the Undulated short-tail (Pax1un-s) mutant, the complete
absence of the Pax1 locus resulted in persistent notochord, loss of all vertebrae and
intervertebral discs, and severe malformations of the ribs and sternum (Wallin, et al.
1994). The Pax1un-s mouse, which was generated by gene targeting to produce a
defined Pax1 null allele, had similar deficits in the vertebral column including scoliosis,
split vertebrae, abnormalities in the lateral vertebral processes, and loss of
intervertebral disc structures (Wilm, et al. 1998). Pax1 null mice had a mild but fully
penetrant phenotype in the vertebral column as well as deficits in the sternum and
scapula (Wilm, et al. 1998). Additionally, 88% of Pax1 heterozygotes were observed to
have some form of defect in sclerotome-derived structures (Wilm, et al. 1998). Despite
these deficits, Pax1 null mutants were viable (Wilm, et al. 1998).
25
Pax1 and Pax9 have partially redundant roles in sclerotome differentiation with
Pax1 appearing to be more important for vertebral column formation (Peters, et al.
1999). Deletion of Pax9 did not result in major deficits in the vertebral column (Peters, et
al. 1999). While removal of Pax1 resulted in some malformations in the vertebrae and
intervertebral discs, removal of both Pax1 and Pax9 resulted in major deficiencies in the
vertebral column including completely absent vertebral bodies and intervertebral discs
(Peters, et al. 1999). Additionally, Pax1;Pax9 null mutants had deficits in the proximal
ribs (Peters, et al. 1999). Interestingly, while Pax1 and Pax9 removal resulted in
agenesis of the ventral vertebrae (vertebral bodies), the lateral vertebrae formed in the
absence of Pax1 and Pax9. Together, these results demonstrated that the ventral and
lateral vertebral processes have distinct genetic requirements for development. Studies
have shown that the ventromedial sclerotome region gives rise to the vertebral bodies
and annuli fibrosi while the dorsomedial and ventrolateral regions form the spinous
processes and neural arches that form the dorsal enclosure for the spinal cord
(Bruggeman, et al. 2012; Christ, et al. 2004; Wallin, et al. 1994; Leitges, et al. 2000;
Adham, et al. 2005).
Mutations in genes known to function upstream of Pax1 and Pax9 also impair
sclerotome differentiation. Deletion of Pbx1/Pbx2, which regulate Pax1 and Pax9 in the
sclerotome, resulted in vertebral defects (Capellini, et al. 2008). Pbx1 and Pbx2 are
expressed in the notochord and surrounding mesenchyme at E12.5 and are important
for chondrocyte differentiation and axial skeleton patterning (Capellini, et al. 2008;
Selleri, et al. 2001). On a Pbx1 null background, removal of one Pbx2 produced thinner
transverse processes and flattened vertebrae at e13.5 (Capellini, et al. 2008). Impaired
26
Shh signaling can also impact sclerotome maturation as hedgehog signaling is required
for Pax1 induction in the sclerotome (McMahon, et al. 1998; Koseki, et al. 1993; Mo, et
al. 1997; Buttitta, et al. 2003).
Bapx1
After the tendon-producing syndetome splits off from the sclerotome, sustained
expression of Pax genes in the remaining sclerotome ensures the continued
specification of chondroprogenitors. Bapx1 is an important mediator of chondrogenic
differentiation that functions downstream of Pax1 and Pax9 in the sclerotome (Rodrigo,
et al. 2003; Tribioli, et al. 1999). Targeted deletion of Bapx1 resulted in impaired
sclerotome differentiation, dysplastic vertebral columns at birth, and perinatal lethality
(Akazawa, et al. 2000). Bapx1 null mutants had unossified vertebrae with a residual
notochord that failed differentiate into nuclei pulposi (Akazawa, et al. 2000).
As differentiation proceeds, Pax1 downregulation in the sclerotome is
accompanied by up-regulated expression in Sox9 (Takimoto, et al. 2013). Sox9 is a
critical regulator of chondrogenic differentiation. Following Pax1 downregulation, Bapx1
expression is maintained in sclerotome-derivatives by Sox9 to suppress osteogenic
differentiation in favor of a chondrogenic pathway (Yamashita, et al. 2009).
Sox Family of Genes
The intervertebral discs are anchored to adjacent vertebrae by the cartilaginous
endplates, which are structurally contiguous with the fibrocartilaginous annulus fibrosus.
Sox9, which is regarded as the master regulator of chondrogenesis, is a SRY-related
HMG box containing transcription factor expressed in the notochord, sclerotome, and
their derivatives (Bi, et al. 1999; Sugimoto, et al. 2013; Barrionuevo, et al. 2006). Sox9
27
and its homologs Sox5 and Sox6 play critical roles in development and maintenance of
cartilage in the vertebral column. Sox9 is expressed in the notochord and is required for
its maintenance and differentiation (Barrionuevo, et al. 2006).
Sox9 is a critical regulator of many of the steps involved in cartilage
differentiation, including the earliest stages of mesenchymal condensation that occurs in
the somitic mesoderm (Bi, et al. 1999; Akiyama, et al. 2002). Mutations in Sox9 underlie
chondrodysplasias in animals and humans, and homozygous inactivation of Sox9 is
known to result in complete absence of cartilage while Sox9 mis-expression induces
ectopic cartilage formation (Akiyama, et al. 2002; Hargus, et al. 2008; Henry, et al.
2012; Bi, et al. 1999; Takimoto, et al. 2012). In the absence of Sox9, loose
undifferentiated sclerotome fails to condense to form chondrogenic nodules, which
usually occurs between 11.5 and 12.5 dpc in mice (Bi, et al. 1999). In chimeric mice
generated with Sox9 null embryonic stem cells, Sox9 null cells failed to express
chondrogenic markers, were excluded from the vertebral cartilage, and were instead
relegated to the supporting mesenchyme (Bi, et al. 1999).
Sox9 is also a critical regulator of postnatal cartilage maintenance. Conditional
inactivation of Sox9 in postnatal mice resulted in arrested development of growth plate
chondrocytes, proteoglycan loss, and down-regulated expression of many cartilage-
specific genes (Henry, et al. 2012). Sox9 inactivation resulted in kyphosis of the
vertebral column and compressed intervertebral discs, mimicking degenerative changes
seen in humans (Henry, et al. 2012).
Sox9 is also critical for its role in transcriptionally activating Sox5 and Sox6 in the
cartilage (Akiyama, et al. 2002). Sox5 and Sox6 cooperate to stabilize Sox9 binding at
28
its target regulatory sequences, thus enhancing transcription of chondrogenic mediators
(Lefebvre, et al. 1998). Sox9, Sox5, and Sox6 have overlapping and distinct roles in
chondrogenic differentiation (Ikeda, et al. 2005; Smits, et al. 2001; Smits, et al. 2003; de
Crombrugghe, et al. 2000). Like Sox9, Sox5 and Sox6 are expressed in the notochord,
sclerotome, and intervertebral discs. Mice that lacked both Sox5 and Sox6 lacked
notochordal sheaths and showed signs of aberrant cell death in the notochord. As a
result, Sox5;Sox6 null embryos developed vertebral columns that lacked nuclei pulposi
(Smits, et al. 2003).
Scleraxis And TGFβ Signaling
Tendons and ligaments are force-transmitting connective tissues that adjoin bone
to muscle and bone to bone, respectively. Within the vertebral column, tendons and
ligaments anchor the vertebrae to each other and to the musculature. Scleraxis (Scx) is
expressed in embryonic and postnatal tendons and ligaments, and is presently the
earliest known marker of tendon/ligament specification in the somitic mesoderm
(Cserjesi, et al. 1995; Brent, et al. 2003). Scx regulates expression of downstream
tenogenic genes including type I collagen, fibromodulin, decorin, tenomodulin, and
tenascin C (Takimoto, et al. 2012; Shukunami, et al. 2006; Murchison, et al. 2007;
Schneider, et al. 2011). In addition, Scx is expressed in the annulus fibrosis, although its
role in the annulus fibrosus is unknown. The annuli fibrosi develop normally in Scx null
mice (Murchison, et al. 2007).
Mice with targeted deletion of Scx were viable but had acute dorsal flexure of the
forelimbs at birth, indicating defects in the limb tendons (Murchison, et al. 2007). In the
vertebral column, Scx loss affected only a subset of tendons, and did not affect the
29
annuli fibrosi (Murchison, et al. 2007). Although Scx null mice lacked most
tendons/ligaments, including the long tendons that anchor the tail vertebrae to the base
of the tail, some short tendons remained unaffected despite Scx loss (Murchison, et al.
2007). Thus, although Scx is presently the earliest known marker of syndetome
specification in the somitic mesoderm, another unidentified regulator is likely
responsible for activating the tenogenic developmental program in the somitic
mesoderm, upstream of Scx.
Another important mediator of tenogenic differentiation includes the TGFβ
signaling pathway. TGFβ2 and TGFβ3 are expressed in cartilage and tendon
progenitors early in development, and TGFβ signaling is known to regulate the
expression of many tendon-specific genes, including Scleraxis, Tenomodulin, and
Mohawk (Schneider, et al. 2011; Liu, et al. 2015; Pryce, et al. 2009). Deletions and
mutations in TGFβ or its signaling constituents affect vertebral column development and
intervertebral disc morphogenesis (Pryce, et al. 2009; Baffi, et al. 2004; Baffi, et al.
2006). In mice, TGFβ signaling disruption resulted in absent trunk tendons and
ligaments, Scx downregulation, and small, abnormally shaped vertebrae and reduced or
absent intervertebral discs (Pryce, et al. 2009; Baffi, et al. 2004).
Conclusions
A balanced synergism between the vertebrae, discs, and surrounding
tendons/ligaments and musculature contributes to the fluid and painless mobility of the
axial skeleton. The timely orchestration of activating and antagonizing signals between
the notochord, somites, neural tube, and surface ectoderm is critical for the formation of
the vertebral column. Any perturbation to this complex interplay of signals is liable to
30
produce defects with functional consequences. In adulthood, the avascular, hypoxic
environment of the intervertebral discs limits its regenerative potential, rendering the
discs vulnerable to age-related degeneration.
Discogenic back pain can be a chronic and debilitating condition. The lifetime
prevalence for back pain is greater than 80% (Andersson, 1999; Katz, 2006). The
current standard of care for back pain largely focuses on pain management through
analgesics and exercise therapy rather than the reversal of underlying pathology. Even
more aggressive treatments, including disc arthroplasty and lumbar spinal fusion, fail to
address the underlying pathology and are limited in efficacy. A more thorough
appreciation of the genetic mechanisms underlying vertebral column formation, and
degeneration, will be requisite for the development of more effective therapies for the
treatment of disc degeneration and back pain.
31
Table 1-1. Genes Required For Murine Vertebral Column Formation
Gene Mutation Phenotype, References
Foxa1 Null No vertebral column phenotype (Maier, et al. 2013)
Foxa2 Null Early embryonic lethality; no notochord forms, precluding vertebral column formation (Ang and Rossant, 1994)
Foxa2 Conditional inactivation in the notochord
Foxa2 inactivation at e7.5 using ShhCreERT2 produces no vertebral column phenotype (Maier, et al. 2013)
Foxa1/Foxa2 Foxa1 null; Foxa2 conditional inactivation in the notochord
Foxa2 inactivation at e7.5 using ShhCreERT2 on a Foxa1 null background results in abnormal vertebral columns with small, misshapen nuclei pulposi (Maier, et al. 2013)
Gdf-5 Null No vertebral column phenotype (Maier and Harfe, 2011)
Gli2 Gli2zfd Reduced vertebral ossification; reduced or absent discs (Mo, et al. 1997)
Gli3 Gli3XtJ Mild neural arch defects; Gli3XtJ/XtJ phenotype less severe than that in Gli2zfd (Hui and Joyner, 1993; Mo, et al. 1997)
Gli2/Gli3 Gli2zfd; Gli3XtJ
Gli2zfd/zfd; Gli3XtJ/XtJ mice are embryonic lethal by E10.5. Gli2zfd/zfd; Gli3XtJ/+ mice have severe sternal defects and vertebral columns with malformed discs and vertebrae (Mo, et al. 1997)
HIF-1α Conditional inactivation in the notochord
HIF-1α inactivation using Foxa2cre resulted in smaller nuclei pulposi at birth. Postnatally, notochord-derived nuclei pulposi cells undergo massive cell death and are replaced by chondrocyte-like cells of non-notochordal origin (Merceron, et al. 2014)
Nkx3.1 Null No vertebral column phenotype (Tanaka, et al. 2000; Herbrand, et al. 2002)
32
Table 1-1. Continued
Gene Mutation Phenotype, References
Nkx3.2 (Bapx1) Null Mild phenotype; ventromedial vertebral elements (vertebral bodies) affected; lack of ossification; abnormal discs (Akazawa, et al. 2000; Lettice, et al. 2001)
Nkx3.1/Nkx3.2 Nkx3.1 null; Nkx3.2 null
Embryonic lethal by e17.5. Reduced ossification and defects in dorsal vertebral elements (transverse processes); anterior vertebral column more severely affected (Herbrand, et al. 2002)
Noggin Null Perinatal lethality; aberrant presence of non-notochordal cells in the notochord; loss of caudal sclerotome and vertebrae; kinked and shortened tails (McMahon, et al. 1998; Tylzanowski, et al. 2006; Li, et al. 2007)
Noto88, 89, 133 Nottc
NoteGFP Reduced viability. Truncation in tail notochord results in loss of tail vertebrae (Theiler, 1959; Abdelkhalek, et al. 2004; Zizic-Mitrecic, et al. 2010)
Pax1 Pax1un Pax1un-s Pax1un-ex Pax1un-i Pax1null
Vertebral column phenotypes of variable severity. Defects include shortened or kinky tails; scoliosis; loss of vertebrae and intervertebral discs. Pax1un-s produces the most severe phenotype and is regarded as semi-dominant as heterozygotes also exhibit vertebral column defects (Wallin, et al. 1994; Wilm, et al. 1998; Adham, et al. 2005)
Pax9 Null No vertebral column phenotype (Peters, et al. 1998; Peters, et al. 1999)
Pax1/Pax9 Pax1 null; Pax9 null Complete loss of ventral vertebral elements (vertebral bodies) and intervertebral discs (Peters, et al. 1999)
Shh Null Cyclopia; complete agenesis of the vertebral column; rudimentary ribs (Chiang, et al. 1996)
Shh Conditional inactivation in the notochord
Inactivation at E7.5 using ShhCreERT2 resulted in complete absence of discs and vertebrae. Shh inactivation at e11.5 produced a normal vertebral column (Choi and Harfe, 2011)
33
Table 1-1. Continued
Gene Mutation Phenotype, References
Sox5 Null Perinatal lethality; vertebral column formed with mild vertebral ossification delay (Smits, et al. 2001)
Sox6 Null Perinatal lethality; vertebral column formed with mild vertebral ossification delay (Smits, et al. 2001)
Sox5/6 Sox5 null; Sox6 null Embryonic lethal at E16.5 with severe defects in the vertebral column including chondrodysplasia; cartilage deficits; small and malformed or absent nuclei pulposi (Smits, et al. 2001; Smits, et al. 2003)
T-brachyury Null Embryonic lethal by E10.5; loss of trunk and tail level notochord precludes vertebral column formation (Gruneberg, 1958; Wilkinson, et al. 1990; Hermann, 1992; Beddington, et al. 1992)
Tbx18 Null Perinatal lethality with shortened axial skeletons. Defects included kinks in the thoracic vertebral column; malformed, flattened discs and vertebral bodies; expanded pedicles and transverse processes (Bussen, et al. 2004)
Tgfbr2 Conditional inactivation in cartilage
Inactivation in chondrocytes using Col2a-cre produced vertebral column defects including missing or malformed discs; neural arch and transverse process defects; loss of spindle cell shape in the annulus fibrosus (Baffi, et al. 2004; Baffi, et al. 2006)
Uncx4.1 Null Perinatal lethality with defects in the lateral vertebral elements (neural arches; pedicles; transverse process); kink in the thoracic vertebral column (Leitges, et al. 2000)
34
Figure 1-1. Anatomy of the Vertebral Column and Intervertebral Disc. A) The fibrocartilagenous intervertebral discs (IVD) separate and cushion the vertebral bodies (VB) along the length of the vertebral column. B) Each intervertebral disc is comprised of a central, hydrated core called the nucleus pulposus, which is encapsulated by the annulus fibrosus. Each disc is adjoined to the vertebrae by the cartilaginous endplates, which serve as the interface through which nutrient diffusion to the discs occurs.
Figure 1-2. Vertebral Column Development In Mice. The vertebral column develops rostrocaudally with overt chondrogenesis, defined as the accumulation of cartilage matrix molecules in the sclerotome, beginning at 12.5dpc in mice. These chondrogenic condensations can be observed as Alcian blue staining in the vertebral column surrounding the notochord, N, at 12.5 dpc. As development progresses, these chondrogenic condensations exert an inward pressure against the notochordal sheath at repeating intervals. These repeating intervals of compression promote the notochord to disc transition, which can be observed by 14.5 dpc. By 15.5 dpc in the rostral embryo, the discs have formed as distinct, repeating units in the vertebral column. By P0 the discs have acquired their characteristic flattened form. Postnatal vertebral column
35
development entails growth and elongation of the vertebral bones by endochondral ossification and proteoglycan accumulation and eventual cell senescence in the discs.
Figure 1-3. Postnatal Intervertebral Disc Development. By birth, the nucleus pulposus is distinct from the annulus fibrosus, which encapsulates the water-rich disc core. Postnatally as the vertebral column grows in size, the discs enlarge largely by extracellular matrix deposition. By three weeks old, proliferation has largely ceased in the intervertebral discs (Dahia 2006). Postnatal disc development is characterized by deposition of additional annulus fibrosus layers and the emergence of the cartilaginous endplates (EP) as a distinct structure at the disc-vertebra interface. As can be seen in postnatal day 21 (P21) discs, the endplates are contiguous with the inner annuli fibrosi. At P21 the vertebrae continue to grow by endochondral ossification, whereby vertebral growth plate chondrocytes (VG) undergo a sequential differentiation process to form new bone. Adult discs, by comparison, are much less cellular than perinatal discs and P21 discs. As can be seen in discs from 3 month old mice, the discs are much less cellular and rich in extracellular proteoglycans. All images are of frontally sectioned, Alcian blue and Picrosirius red stained discs from wildtype mice.
36
CHAPTER 2 ROLE OF ROBO GENES DURING MURINE VERTEBRAL COLUMN DEVELOPMENT
Introduction
The vertebral column is comprised of ossified vertebrae which are adjoined by
the cartilaginous intervertebral discs. In adult animals, the intervertebral discs can be
characterized as having three functionally distinct domains. Each intervertebral disc is
comprised of a gel-like inner core called the nucleus pulposus (Choi and Harfe, 2011).
The nuclei pulposi are encapsulated by the fibrocartilagenous annuli fibrosi, which can
be further divided/specified/distinguished as inner and outer annuli fibrosi based on
extracellular matrix (ECM) composition (Cortes, et al., 2012). Contiguous with the annuli
fibrosi are the cartilaginous endplates, which serve as the interface between the discs
and the vertebral growth plates (Maladrino, et al., 2014). In adulthood, the discs are
aneural and avascular, leading to a hypoxic environment for nucleus pulposus cells
(Richardson, et al. 2012). Additionally, as growth occurs and the vertebral column
enlarges, nutrient supply to the discs becomes limited to diffusion through the
endplates, which can become calcified and pathologic with age. Altogether, these
changes are thought to contribute to intervertebral disc pathology and back pain in
humans (Tolofari, et al., 2010; Smith, et al., 2011).
The intervertebral discs form from the axial mesoderm (notochord) and paraxial
mesoderm (somites) while the vertebrae form from the paraxial mesoderm (Christ and
Scaal, 2008; Christ, et al., 2004; Choi and Harfe, 2011). Within the paraxial mesoderm,
progenitor cells in the sclerotome give rise to the vertebrae and fibrocartilagenous
components of the discs (endplates and annuli fibrosi) (Bruggeman, et al., 2012; Christ
37
and Scaal, 2008). The nuclei pulposi are known to derive exclusively from the
embryonic notochord (Choi and Harfe, 2011; McCann, et al. 2012).
During vertebral column morphogenesis, cells in the sclerotome differentiate and
form chondrogenic condensations. In mice overt chondrogenesis, defined as the
accumulation of Col2a1 in these condensed regions, beings at approximately 12.5 dpc
in the rostral embryo. As these mesenchymal nodules condense against the
notochordal sheath, notochordal cells are pushed into nucleus pulposus anlagen,
resulting in the formation of vertebrae and intervertebral discs in tandem (Choi and
Harfe, 2011).
Robo proteins are transmembrane cell surface receptors which belong to the
immunoglobulin class of cell adhesion molecules (Blockus and Chedotal, 2016). Four
Robo receptors have been identified and characterized in mice. Robo receptors have
extracellular domains consisting of three fibronectin type III domains and five Ig-like
domains, which interact with secreted glyocoprotein ligands (Hohenester, 2008). The
primary ligands for Robo receptors are Slit proteins (Hohenester, 2008). Heparin
sulfates serve as co-receptors to stabilize Robo-Slit interactions. Three Slit genes have
been identified in mice. Slit proteins undergo proteolytic processing to produce cleavage
fragments that have unique functional properties (Ordan, et al., 2015). Robo receptors
also have conserved intracellular domains that are important for transducing Slit-Robo
signals to intracellular proteins (Ypsilanti, et al., 2010). These conserved domains
interact with cytoplasmic adaptor proteins, including srGAP proteins, or Slit-Robo
GTPase activating proteins, which mediate the effects of Slit-Robo signaling (Ypsilanti,
et al., 2010).
38
The term „promiscuous binding‟ has been used to describe Slit-Robo interactions,
reflecting the ability of any Slit protein to bind to any of the four Robo receptors
(Ypsilanti, et al., 2010). Additionally, Robo receptors are predicted to interact with other
proteins belonging to the LRR family of proteins, which includes the Slit proteins (Howitt,
et al., 2004). Examples of other small LRR proteins include fibromodulin, decorin, and
biglycan, which are expressed in cartilaginous tissues including the intervertebral discs
(Chen, et al., 2015).
In vitro and in vivo studies have shown that Slit-Robo interactions mediate a
chemorepulsive response in Robo-expressing cells (Domyan, et al., 2014; Kim, et al.,
2015). Unlike other signaling pairs like Ephrin-Eph receptor interactions, which result in
bi-directional signaling, Slit-Robo interactions are not known to elicit a response in Slit-
expressing cells. Thus, Slit-Robo signaling is unidirectional. Additionally, Slit-Robo
signaling is known to stimulate changes in cell polarity, adhesion, and migration by
modulating the localization of microtubule organizing centers (MTOC). These changes
are mediated through modulation of cytoskeletal dynamics in a RhoA-dependent
manner (Ypsilanti, et al., 2010).
A role for Robo and Slit in chondrocyte differentiation and function can be
inferred from reports of Robo and Slit expression in chondrocytes. In chicks and rats, for
example, Robo and Slit mRNA are expressed by limb bud chondrocytes in the
perichondrium and appendicular growth plates (Noel, et al., 1998; Holmes and
Niswander, 2001). In rat tibial bones, for example, Robo genes are expressed in the
growth plates (Noel, et al., 1998). Additionally, Robo mRNA expression can be induced
39
by mechanical loading, suggesting a role for Robo receptors in cartilage and bone
remodeling (Noel, et al., 1998)
At present, few studies have investigated the role of Robo and Slit genes in the
sclerotome or its derivatives. We found that in mice, Robo1 and Robo2 were expressed
in the sclerotome and had complementary expression patterns to Slit mRNA in the
notochord. In the absence of Robo1 and Robo2, proliferation in the sclerotome was
impaired and resulted in malformed intervertebral discs. Additionally, Robo1 and Robo2
loss in the sclerotome resulted in deficits in the distal ribs with accompanying defects in
the sternum.
Results
Complementary Robo and Slit Expression Patterns in the Vertebral Column Robo Expression
To determine where and when Robo and Slit are expressed during formation of
the vertebral column we used RNA in situ hybridization, antibodies, and Xgal staining.
LacZ knockin into the Robo1 or Robo2 loci was used to generate the Robo1 and Robo2
null alleles. Robo expression was tracked by performing Xgal staining in Robo1+/-
;Robo2+/- mice (also referred to as Robo1;Robo2 heterozygotes in this manuscript). For
Xgal staining, embryos were whole mount stained and then sectioned for imaging by
bright field microscopy. For RNA in situ hybridization, probe templates for Robo1 and
Robo2 were generously provided by Dr. Xin Sun.
Robo expression in the mouse neural tube has been reported previously
(Camurri, Sundaresan, et al, 2004). To confirm specificity of our Robo probes, wildtype
10.5dpc embryos were used for in situ hybridizations. Robo2 mRNA was identified in
40
the neural tube by RNA in situ hybridization (Fig. 2-1A). To confirm these results, stage-
matched Robo1;Robo2 heterozygous embryos were Xgal stained. Robo expression in
the neural tube at 10.5 dpc was confirmed by Xgal staining (Fig. 2-1C). Similarly, Robo2
in situ hybridization and Xgal staining yielded congruent results for Robo expression in
the vertebral column at 12.5 dpc (Fig. 2-1B and Fig. 2-1D). Thus, Xgal staining is a
reliable method to show Robo1 and Robo2 gene expression in the vertebral column.
At 12.5dpc Robo expression was identified in the somitic mesoderm, both lateral
and ventral to the notochord (Fig. 2-2A-B). By RNA in situ hybridization, Robo1 and
Robo2 were detected in the somitic mesoderm of the vertebral column (Fig. 2-2C-D).
Notably, while Robo2 is robustly expressed at the midline, dorsal and ventral to the
notochord (Fig. 2-2D), Robo1 mRNA was not detected at the midline (not shown).
Instead, a repeating pattern of Robo1 expression was observed in the lateral vertebral
column mesenchyme, laterally adjacent to the notochord (Fig. 2-2C). At 14.5 dpc,
Robo1 and Robo2 were expressed in the presumptive annuli fibrosi, which
encapsulated the developing nuclei pulposi (Fig. 2-2E-F). At 18.5 dpc, Robo2 mRNA
was expressed in the spinal cord and in the vertebral column, albeit more diffusely than
at earlier stages (Fig. 2-2G).
Following whole mount Xgal staining of Robo1;Robo2 heterozygotes, a
microtome was used to generate 10um cross-sections of the vertebral column. Using
this method, Robo1 and Robo2 expression was observed in the vertebral trabeculae
(Fig. 2-2I) and in isolated annulus fibrosus cells (Fig. 2-2J) at P0. We did not detect
Robo expression in the nucleus pulposus at any time. To generate a more global
impression of Robo expression in the vertebral column, we used agarose-embedded,
41
Xgal-stained tissues to generate 100um sections on a vibratome. Using this method
Robo1 and Robo2 expression was found to be enriched in the vertebral growth plates
(Fig. 2-2H).
Slit Expression
A number of studies have reported Slit expression in the embryonic floor plate
and notochord3. RNA in situ hybridization confirmed Slit1 and Slit2 expression in the
mouse floor plate and notochord at 10.5 dpc (Fig. 2-3A-B) and 12.5 dpc (Fig. 2-3D-E).
Slit3 mRNA was also detected in the floor plate at 10.5 dpc and 12.5 dpc (Fig. 2-3C and
Fig. 2-3F). At 15.5 dpc, Slit2 expression was maintained and enriched in the nuclei
pulposi (Fig. 2-3H). Slit1 and Slit3 were not expressed at appreciable levels in the nuclei
pulposi of wildtype 15.5 dpc embryos (Fig. 2-3G and Fig. 2-3I).
Robo and Slit maintained complementary expression patterns during vertebral
column morphogenesis. At 12.5 dpc, for example, Robo1 and Robo2 were expressed in
the somitic mesoderm lateral and ventral to the notochord, but were not expressed in
the notochord (Fig. 2-2A-B). By comparison, Slit2 mRNA was expressed in the
notochord but not in the surrounding mesenchyme (Fig. 2-3E). This complementarity
was maintained until at least 14.5 dpc, after initiation of the notochord to nucleus
pulposus transition. At 14.5 dpc Robo1 and Robo2 were expressed in the presumptive
annuli fibrosi (Fig. 2-2E-F) while in comparably aged embryos, Slit2 mRNA was
enriched in the nuclei pulposi anlagen (Fig. 2-3H). Thus, Robo and Slit exhibit
complementary expression patterns during vertebral column morphogenesis in mice.
42
Intervertebral Disc Malformations
The Robo1 and Robo2 null alleles, which reside 1.1 cM apart on chromosome 16
in mice, were previously generated by LacZ knockin to investigate their role in kidney
development (Long, et al, 2004; Grieshammer, et al, 2004). Robo1;Robo2 null mice
were then generated by homologous recombination of the Robo1 and Robo2 null alleles
to produce linked mutant alleles (Domyan, et al, 2013). As described previously,
Robo1;Robo2 null mice die immediately after birth due to respiratory defects but are
comparable in size to control littermates at birth (Domyan et al, 2013).
Alcian blue and Picrosirius red were used to histologically examine the effects of
Robo1 and Robo2 removal vertebral column morphogenesis at P0. In control mice the
vertebral column was comprised of repeating units of ossified vertebrae separated by
characteristically flattened intervertebral discs. By comparison, the intervertebral discs
in Robo1;Robo2 null mice were bulbous in appearance and had reduced Alcian blue
staining (Fig. 2-4A).
Homozygous removal of Robo1 alone did not produce a defect in the
intervertebral discs (Fig. 2-4B). To assess the potential role of Slit proteins in mediating
the Robo-dependent phenotype, Slit1;Slit2 null mutants were examined. Intervertebral
discs from Slit1;Slit2 null mice appeared comparable to those from control littermates at
17.5 dpc (Fig. 2-4C). Together, these data suggest a functional redundancy between
Robo1 and Robo2 in intervertebral disc development and show that removal of Slit1 and
Slit2 does not recapitulate the Robo1;Robo2 null phenotype in the vertebral column.
To confirm that Robo1 and Robo2 loss in the somitic mesoderm was the cause
of the chondrogenic deficits and intervertebral disc malformations, Robo2 was
43
conditionally inactivated on a Robo1 null background using a Robo2Floxed allele.
Inactivation of Robo2Floxed using ShhCre resulted in a normal vertebral column at P0
(Fig. 2-5C). Inactivation of Robo2Floxed using Dermo1Cre partially recapitulated the
Robo1;Robo2 null phenotype (Fig. 2-5B). Compared to control littermates, Robo1-/-
;Robo2F/F;Dermo1Cre mice had reduced Alcian blue staining, suggesting a deficit in
cartilage formation or maintenance following formation of the vertebrae and discs (Fig.
2-5B).
The Notochord to Nucleus Pulposus Transition
One of the hallmarks of disc degeneration is fibrogenesis and eventual loss of
„notochord-like‟ cells in the nucleus pulposus (Lv, Peng, et al, 2016). In mice, for
example, removal of HIF1a in the discs resulted in complete loss of normal nucleus
pulposus cells by apoptosis, and eventual replacement of those cells by fibroblast-like
cells of unknown origin. These changes were accompanied by deficits in vertebral
column mechanical function. Moreover, one of the characteristics of aging discs in
humans is the gradual loss or effacement of the nucleus pulposus-annulus fibrosus
boundary (Urban and Roberts, 2003). Thus, normal disc biology entails strict
segregation of nucleus pulposus cells from annulus fibrosus cells during development
and loss of this separation is associated with disc degeneration.
Fate mapping studies in mice have shown that adult nuclei pulposi are comprised
of cells exclusively of notochordal origin. Equally, cells of non-notochordal origin are
excluded from (or undergo apoptosis) the inner nucleus pulposus in adult mice,
although the mechanism underlying this phenomenon is unknown.
44
In other developing tissues, Robo-Slit signaling is essential for modulating cell
migration and organ positioning via a chemorepulsive mechanism. In the developing
central nervous system (CNS), for example, Robo-expressing commissural axons are
repelled from the midline by responding to chemorepulsive Slit cues secreted from the
neural tube floor plate. In the CNS, Slit-mediated repulsion of Robo-expressing
commissural axons prevents aberrant re-crossing of axons across the midline. In the
developing foregut, Robo-expressing foregut mesenchyme responds to secreted Slit
ligands from the body wall to ensure that the developing gut becomes properly
positioned in the abdominal cavity.
The observation that Slit and Robo have complementary expression patterns
during vertebral column formation (Fig. 2-2 and Fig. 2-3) suggested a potential role for
Robo-Slit signaling in mediating the notochord to nucleus pulposus transition in mice.
The expression patterns of Slit and Robo and their known functions in other tissues
suggested a model in which Robo-expressing cells were repelled by Slit expression in
the notochord and nuclei pulposi (Fig. 2-6A).
In this model, Slit-mediated repulsion of Robo-expressing cells would prevent
mixing of nucleus pulposus and annulus fibrosus cells during vertebral column
morphogenesis (Fig. 2-6A). Based on this model, removal of Robo1 and Robo2 would
impair the inhibitory effect of Slit expression in the nucleus pulposus on annulus fibrosus
cells, resulting in a mixed population of cells in the nuclei pulposi at birth (Fig. 2-6A).
Specifically, this mixture of cells would include cells of notochordal origin and cells of
non-notochordal (sclerotomal) origin. To address this possibility, and to determine the
45
developmental origin of nucleus pulposus cells at birth, the Rosa-mT/mG dual
fluorescence reporter was used.
The Rosa-mT/mG reporter was generated by insertion of the mT/mG construct at
the constitutively active Rosa26 locus (Muzumdar, et al., 2007). In the absence of Cre,
mice that carry the Rosa-mT/mG allele have membrane-targeted tdTomato
fluorescence in all cells of the intervertebral discs. In the presence of Cre, excision of
the tdTomato cassette via flanking loxP sites results in expression of membrane-
targeted GFP in lieu of membrane-targeted tdTomato. Lineage tracing analyses by Cre-
mediated homologous recombination results in the „genetic marking‟ of all Cre-
expressing cells as well as all daughter cells derived from the Cre-expressing cell
population. In these experiments activation of GFP is permanent such that all daughter
cells are irreversibly GFP-marked, even in the absence of active Cre expression.
Shh is expressed in the nascent notochord and continues to be expressed as the
notochord begins to transition into the nuclei pulposi (Choi and Harfe, 2011). A ShhCre
allele was used to genetically mark the notochord and its derivatives, the nucleus
pulposus. Previous studies have reported a small number of notochord-derived cells
scattered throughout the vertebral column outside of the nuclei pulposi. These cells are
“notochordal remnants” and are potentially the cell population that gives rise to
chordomas. The majority of notochord cells form nucleus pulposus cells with only a few
notochord-derived cells residing outside the nuclei pulposi. Additionally, under normal
circumstances, sclerotome-derived cells are strictly excluded from the nucleus pulposus
(Harfe and Choi, 2011).
46
To assess if Robo-Slit signaling was responsible for maintaining separation of
nucleus pulposus (NP) and annulus fibrosus (AF) cells in the discs, Robo1-/-; Robo2-/-;
ShhCre; Rosa-mT/mG mice were generated and harvested at 18.5 dpc for lineage
tracing analyses (Fig. 2-6B). In the absence of Robo1 and Robo2, the notochord to
nucleus pulposus transition occurred normally (Fig. 2-6C). Notochord-derived cells were
found largely within the nuclei pulposi of Robo1;Robo2 null mice (Fig. 2-6C). Both
controls and Robo1;Robo2 null mice had isolated GFP+ cells in the vertebral column
outside of the nuclei pulposi (not shown). In addition, non-notochord-derived tdTomato+
cells were completely excluded from the nuclei pulposi in both control and
Robo1;Robo2 null mutants (Fig. 2-6C). These data indicate that in the absence of
Robo1 and Robo2, the notochord to nucleus pulposus transition occurs normally.
Using the ShhCre allele, it is impossible to determine if an annulus fibrosus
progenitor cell aberrantly migrated into the nucleus pulposus and began to express Shh
as a consequence of its ectopic location in the notochord. If this occurred, the ectopic
cell would express Shh and activate GFP, making it indistinguishable from Shh-
expressing notochord-derived cells located in the nucleus pulposus. Notochordal cell-
conditioned media has been shown to modulate annulus fibrosus cell gene expression
in vitro. In addition, pluripotent cells are known to acquire notochord-like cell phenotypes
when cultured on nucleus pulposus tissue matrix. These data suggest that other cell
types, if located in the nucleus pulposus, could assume a nucleus pulposus cell-like
expression profile.
To address the limitations of using an approach that could not discriminate
nucleus pulposus cells from cells that had inappropriately migrated and assumed a
47
nucleus pulposus-like expression profile, the lineage tracing analyses were repeated
using the Tamoxifen-inducible ShhCreERT2 allele. In ShhCreERT2 mice, Cre remains
cytoplasmic and inactive in the absence of Tamoxifen. Upon Tamoxifen treatment, Cre
translocates into the nucleus and mediates homologous recombination of DNA flanked
by loxP sites.
Previous data indicates that Tamoxifen administered by oral gavage genetically
marks cells over a 24 hour window in most tissues including the discs (Harfe and Choi,
2011). Using a single dose of Tamoxifen administered to pregnant dams at 10.5 dpc
specifically marks Shh-expressing cells between 10.5 dpc and 11.5 dpc. In the
developing vertebral column, these experiments labeled all cells of the notochord (Harfe
Choi, 2011). By the time the nucleus pulposus formed at ~13.5 dpc, any non-
notochordal cells located in the nucleus pulposus that expressed Shh would not be
marked (ie. GFP-positive). For example, ectopic annulus fibrosus or end plate cells that
inappropriately migrated into the notochord after 11.5 dpc would not be expected to be
GFP-positive even if they were located in the nucleus pulposus and expressed Shh.
Using the above strategy, no mixing of cells in the nucleus pulposus was
observed in Robo1;Robo2 null mutants. All nucleus pulposus cells were GFP-positive
and all tdTomato-positive cells were strictly excluded from the nucleus pulposus region
in both control and Robo1;Robo2 null animals (Fig. 2-7B). A notable difference between
control and Robo1;Robo2 null mice was that a greater number of GFP+ cells were
observed outside the nuclei pulposi in Robo1;Robo2 mutants (Fig. 2-7B). Although a
few isolated GFP+ cells were observed outside the nuclei pulposi in controls (not
48
shown), Robo1;Robo2 null mice had a greater number of GFP+ cells in the annuli fibrosi
and vertebrae (Fig. 2-7B).
These data can be interpreted in at least two ways. First, it is possible that GFP+
cells located outside the nuclei pulposi in Robo1;Robo2 null mice are notochordal in
origin. It is possible that in the absence of Robo1 and Robo2, the notochord to disc
transition is affected and notochord-derived cells become ectopically scattered
throughout the vertebral column.
Alternatively, it is possible that the GFP+ cells outside the nuclei pulposi are
sclerotomal in origin. A Tamoxifen pulse at 10.5dpc would mark all cells expressing Shh
between 10.5 and 11.5dpc, including cells in the notochord. This would also mark
isolated progenitors in the sclerotome that may have transiently expressed Shh during
the same window. Shh expression and function in the sclerotome has not been reported
previously.
One way to resolve this uncertainty would be to co-label GFP+ cells from Rosa-
mT/mG mice with an anti-T-brachyury antibody. T-brachyury is expressed in the
notochord and its expression, as determined by T-brachyury protein
immunofluorescence, is maintained in the nucleus pulposus at P0. If GFP+ cells outside
the nuclei pulposi co-labeled with T-brachyury, it would suggest that these GFP+ cells
were notochord-derived.
Rib and Sternal Development
In addition to abnormalities in the vertebral column, deficits in the distal ribs and
sterna were observed in Robo1;Robo2 null mice. Rib cages from Robo1;Robo2 null
mice were smaller in comparison to those from control littermates (Fig. 2-8A). Closer
49
examination revealed that in addition to having smaller rib cages, Robo1;Robo2 null
mice had deficits in the distal ribs and sterna (Fig. 2-8B). Specifically, compared to
controls which had attachment of the cartilaginous distal ribs at the sternum, (Fig. 2-8B,
a-b), the distal ribs in Robo1;Robo2 null mice failed to meet at the midline (Fig. 2-8B, b-
f). Additionally, the distal ribs showed reduced Alcian blue staining, suggesting a
decrease in chondrogenesis in the distal ribs (Fig. 2-8B, e-f). Moreover, while the
sterna in control animals was always completely fused at birth (Fig. 2-8B, a-b),
Robo1;Robo2 null mice had incomplete fusion of the sternal bars at the midline (Fig. 2-
8B, c-f). To quantify these deficits, sternal length was measured. At P0, Robo1;Robo2
null mice had a 30% reduction in sternal length compared to controls (Fig. 2-8C).
Proliferative Deficits in the Sclerotome
Based on observations that the ribs and sterna were reduced in size in
Robo1;Robo2 null mice, we speculated that Robo1;Robo2 null mice had a proliferative
deficit earlier in development. The ribs and sternum, as well as the vertebrae and annuli
fibrosi, are derived from the sclerotome.
To address the possibility that the ribs and sterna in Robo1;Robo2 null mice were
smaller due to proliferative differences in the sclerotome, an EdU-based assay was
used to assess in vivo proliferation at 13.5 dpc. EdU is a thymidine analogue that
becomes incorporated into newly synthesized DNA as cells undergo mitosis. Using a
fluorescent antibody to detect incorporated EdU, the effect of Robo1 and Robo2 loss on
proliferation was quantified.
Although Robo1 and Robo2 antibodies are commercially available, we were
unable to detect Robo using these antibodies, even in positive control brain tissues. In
50
lieu of direct detection of Robo protein, an antibody against β-Galactosidase (bGal)
protein, which is produced from the lacZ transcript transcribed from the Robo1 and
Robo2 loci, was used. bGal immunofluorescence recapitulated the Robo expression
pattern as visualized by Xgal staining (Fig. 2-9A).
To assess differences in proliferation, we used Robo1;Robo2 null mutants and
Robo1+/-;Robo2+/- heterozygous control littermates. Following the manufacturer‟s
protocol for EdU detection (Fig. 2-9B, d), tissues were incubated with an anti-bGal
primary antibody, followed by incubation with a 488-conjugated secondary, to mark
Robo expression domains in the vertebral column (Fig. 2-9B, b).
Compared to control littermates, Robo1;Robo2 null embryos exhibited decreased
proliferation. Removal of Robo1 and Robo2 resulted in a greater than 50% reduction in
sclerotomal proliferation at 13.5dpc (Fig. 2-9C). Approximately 26% of cells in the
sclerotome were found to be proliferative in controls while only 11% of cells in the
sclerotome were proliferative in Robo1;Robo2 mutants (Fig. 2-9C, d). To determine if
the proliferative deficit was due to an overall decrease in mitotic index caused by Robo
loss, proliferation in the notochord, which does not express Robo1 or Robo2, was
quantified. No difference in proliferation was observed in the notochord (Fig. 2-9C, d).
Robo and Pax1 Expression in the Vertebral Column
Many of the molecular regulators of sclerotome induction and differentiation have
been identified, including proteins belonging to the Meox, Bapx, and Pax family of
transcription factors. One of the earliest markers of sclerotome specification in the
paraxial mesoderm is Pax1. Pax1 is required for sclerotome specification and
differentiation, and in its absence rib and vertebral defects occur. Midline-derived
51
Hedgehog signals are required for Pax1 induction in the paraxial mesoderm. Once
activated, Pax1 directly regulates Bapx1 expression, which is required for subsequent
chondrogenic differentiation of sclerotome-derived tissues.
Pax1 mutants have been reported to have rib and vertebral defects which are
reminiscent of the deficits observed in P0 Robo1;Robo2 null mice (Fig. 2-8A).
Additionally, like Robo1;Robo2 null mice which showed decreased proliferation (Fig. 2-
9C), Pax1 removal has been shown to cause decreased cell proliferation in the
sclerotome (Peters, et al., 1999). To investigate potential interactions between Pax1,
Robo1, and Robo2, we characterized Pax1, Robo1, and Robo2 expression during the
early stages of vertebral column morphogenesis.
At 12.5dpc, Pax1 protein and Robo1 and Robo2 were expressed in the somitic
mesoderm in spatially similar but not identical domains (Fig. 2-10A, a-b). Specifically,
Robo1 and Robo2 expression appeared to be confined to the more medial
mesenchyme closer to the notochord (Fig. 2-10A, a), while Pax1 expression extended
to the lateral mesenchyme (Fig. 2-10A, b). Pax1 expression also appeared broader in
the lateral mesenchyme (Fig. 2-10A, b). Similarly, in the caudal region of 13.5 dpc
embryos Robo2 mRNA and Pax1 protein were expressed in a similar metameric pattern
in the dorsal and ventral mesenchyme surrounding the notochord (Fig. 2-10A, c-d).
Finally, at 14.5 dpc as the notochord to nucleus pulposus transition occurred, Robo1
and Robo2 and Pax1 shared a similar fan-shaped expression pattern in the presumptive
annuli fibrosi (Fig. 2-10A, e-f).
These data suggest that Robo1 and Robo2 are expressed in the embryonic
sclerotome, which is molecularly distinguishable within the somitic mesoderm by Pax1
52
expression. To establish whether Robo1 and Robo2 were co-expressed with Pax1 in
the same cells, vertebral columns from 13.5 dpc embryos were incubated with
antibodies to Pax1 and bGal for confocal microscopy analysis. Confocal microscopy
revealed that although bGal and Pax1 had similar expression domains in the vertebral
column, Pax1-bGal co-localization, as defined as yellow fluorescence that occurs when
Pax1-555 and bGal-488 are in close proximity, occurred in only a subset of cells (Fig. 2-
10B).
Robo Gene Expression and Function in the Growth Plates
The bony vertebrae develop by endochondral ossification, wherein an avascular
cartilaginous template is first established by progenitors in the sclerotome. Following
formation of a cartilaginous skeletal element, cartilage-forming chondrocytes are
eventually replaced by bone forming osteoblasts as the cartilage template undergoes
remodeling and vascularization in preparation for ossification. By P0, the vertebrae have
begun to ossify and chondrocytes become organized at the vertebral growth plates, as
evidenced by the expression of distinct molecular markers. Bone elongation via
endochondral ossification occurs as growth plate chondrocytes, organized into distinct
layers, undergo stages of chondrocyte maturation. The coordinated differentiation of
chondrocytes at the rostal and caudal ends of the vertebra results in appositional growth
and elongation of the vertebrae.
Chondrocytes in the vertebral growth plates can be distinguished as being part of
the quiescent, resting zone (RZ); the proliferative, pre-hypertrophic zone (PZ); the
hypertrophic zone (HZ); and finally, as terminally differentiated, apoptotic cells that give
way to ossification (Fig. 2-11A). Growth plate chondrocytes can be distinguished based
53
on cell morphology (ie. enlarged, hypertrophic cells) and by expression of specific
molecular markers. Chondrocyte differentiation and progression through these growth
plate zones is tightly regulated by a complex interaction of many regulatory factors,
including Bapx1, Pax1, Sox9, Runx2, PTHrP, and Ihh.
During bone growth, quiescent cells in the resting zone (RZ) express PTHrP.
PTHrP is a secreted signaling protein that promotes proliferation and inhibits terminal
differentiation via interaction with the PTH receptor. In response to PTHrP, cells
adjacent to the resting zone become proliferative and begin to express Ihh. Ihh, which is
expressed by pre-hypertrophic cells, feeds back to resting zone cells to stimulate PTHrP
secretion. Thus, the Ihh-PTHrP feedback loop is important for the maintenance of
chondrocytes in an immature state of differentiation. As bone elongation occurs and the
available amount of PTHrP diminishes, cells further away from the resting zone
differentiate further and become hypertrophic. Hypertrophic chondrocytes express
Col10a.
Chondrocytes at the disc-vertebra junction of control animals appeared
organized with distinct layers of small, flattened chondrocytes near the discs, and layers
of larger hypertrophic chondrocytes nearer to the trabecular centers of the vertebrae
(Fig. 2-11A, b) In contrast, chondrocytes at the disc-vertebra junction in Robo1;Robo2
null mice appeared disorganized (Fig. 2-11A, d). In P0 vertebral columns Robo1 and
Robo2 are expressed in the vertebral growth plates (Fig. 2-11B). Robo1 and Robo2
removal resulted in decreased Ihh and Col10a expression (Fig. 2-11C). While Ihh and
Col10a were expressed as distinct, tight bands along the vertebral growth plates in
controls, Ihh in Robo1;Robo2 null growth plates appeared diffuse (Fig. 2-11C, b) and
54
Col10a expression was markedly diminished in mutants (Fig. 2-11C, d). Together, these
data suggest a role for Robo1 and Robo2 in regulating growth plate chondrocyte
differentiation.
Annulus Fibrosus Cell Morphology and Gene Expression
Under normal circumstances, Ihh and Col10a1 expression is restricted to growth
plate chondrocytes and is not expressed in the intervertebral discs (Fig. 2-12B, a, c). In
Robo1;Robo2 null mice, however, ectopic Ihh and Col10a expression was observed in
annulus fibrosus cells (Fig. 2-12B, b and d). These changes were accompanied by cell
shape changes in Robo1;Robo2 null annuli fibrosi, which can be observed by histology
(Fig. 2-12A) as well as by membrane-targeted fluorescence in Rosa-mT/mG mice (Fig.
2-6 C).
Cartilage Matrix Protein Expression
Based on observations that Robo1;Robo2 null mice had decreased Alcian blue
staining in the intervertebral discs (Fig. 2-4), antibodies specific for cartilage matrix
proteins, including Col6a1, Col2a1, Tenascin C, and Aggrecan, were used to assess
potential differences in matrix composition by immunofluorescence. Col2a1, Col6a1,
and Aggrecan are critically important components of cartilage. Col2a1 and Aggrecan
loss is associated with intervertebral disc pathology in humans.
By immunofluorescence a qualitative decrease in Col6a1 expression was
observed in the annuli fibrosi of Robo1;Robo2 null discs (Fig. 2-13A). Col2a1
expression appeared comparable, and was enriched in the inner annuli fibrosi of both
controls and mutants (Fig. 2-13B). Aggrecan immunofluorescence also appeared
diminished in Robo1;Robo2 null tissues (Fig. 2-13C). Aggrecan appeared reduced in
55
both the annuli fibrosi and nuclei pulposi in Robo1;Robo2 null mice (Fig. 2-13C). A
decrease in Aggrecan in the vertebral column was observed in Robo1;Robo2 null mice
as early as 16.5 dpc (not shown).
Tenascin C Upregulation
Tenascin C (TNC) is associated with cartilaginous, ligamentous, and tendinous
tissues and its expression can be induced by mechanical stimulation. In the
intervertebral discs TNC is enriched in the annulus fibrosus. TNC is transcriptionally
regulated by PRX1, and is downstream of the PDGF and TGFbeta signaling pathways.
TNC null mice exhibit deficits in wound healing, but the in vivo consequence of TNC
knockout on chondrogenic differentiation is unknown. TNC belongs to a class of matrix
molecules known as adhesion modulatory proteins. In vitro, TNC has been shown to
modulate cell shape and adhesion by interfering with cell interactions with fibronectin.
Compared to cells plated on fibronectin alone, which were elongated and adherent,
cells plated on fibronectin and tenascin C detached from the substratum and showed
cell rounding.
Based on immunofluorescence, a decrease in TNC protein was observed in
Robo1;Robo2 null discs (Fig. 2-14A). Quantification of these differences by western blot
analysis revealed that Robo1;Robo2 null discs had a 60% increase in 170kD and 250kD
TNC fragments compared to control discs (Fig. 2-14B-C). An increase in the abundance
of 170kD and 250kD TNC fragments has been linked to joint pathology in humans. In
the same report it was demonstrated that recombinant 170kD and 250kD sized TNC
induced cartilage matrix degradation. These data suggest that Robo receptor proteins
56
may play a role in cartilage matrix homeostasis by a Tenascin C-dependent
mechanism.
Discussion and Conclusions
In our studies the combined role of Robo1 and Robo2 in vertebral column
development was investigated using mice that lacked Robo1 and Robo2. The Robo1
and Robo2 individual null alleles were previously generated by LacZ knock-in and the
Robo1;Robo2 null mouse was subsequently generated by homologous recombination
to produce linked mutant alleles (Grieshammer, et al, 2004; Long, et al, 2004; Domyan,
et al, 2013). Robo1 and Robo2 are located 1.1 cM apart on chromosome 16 in mice. As
described previously, removal of Robo1 and Robo2 resulted in perinatal lethality caused
by respiratory defects (Domyan, et al, 2013). As such, previous studies were limited to
investigation of Robo1 and Robo2 expression and function only during embryonic
stages.
Role of Robo1 and Robo2 in IVD Development
Mice that lacked both Robo1 and Robo2 were observed to have intervertebral
disc defects (IVD). Macroscopically, Robo1;Robo2 null discs were bulbous rather than
flattened and disc-shaped as in controls (Fig. 2-4). Additionally, cells in the annulus
fibrosus lacked their characteristic spindle-shaped morphology and lamellar
organization around the nucleus pulposus (Fig. 2-4; Fig. 2-12). Robo1 and Robo2 were
expressed in the somitic mesoderm at 12.5 dpc and were distinctly absent in the
notochord (Fig. 2-2). More specifically, co-localization experiments showed that Robo1
and Robo2 expression domains overlapped with that of Pax1, which is a molecular
marker for sclerotome specification (Fig. 2-10). These data suggest that Robo1 and
57
Robo2 were expressed in the sclerotome, which contains annulus fibrosus progenitors.
Robo1 and Robo2 were also observed in the presumptive annuli fibrosi surrounding the
rudimentary nuclei pulposi at 14.5 dpc, suggesting a potential role for Robo receptors in
annulus fibrosus differentiation from the sclerotome (Fig. 2-2).
Lineage tracing studies using genetic methods have shown that in mice, cells in
the notochord and sclerotome remain separate during morphogenesis of the
intervertebral discs. Nuclei pulposi in adult mice were exclusively of notochordal origin.
Equally, almost all notochord-derived cells were excluded or removed from the
vertebrae and annuli fibrosi (Choi and Harfe, 2011). Notochordal cells that remained in
the vertebrae were postulated to be notochordal remnants, which are thought to be the
cell reservoir responsible for chordoma formation in humans. The mechanism that
ensures proper separation of notochordal and sclerotomal cell populations during the
formation of the intervertebral discs is unknown.
In mice it has been shown that removal of Sox5 and Sox6, or inactivation of the
Hedgehog signaling pathway, results in aberrant scattering of notochord-derived cells in
the vertebrae and annuli fibrosi. Consequently, Sox5;Sox6 null and Hedgehog signaling
mutants had incompletely formed vertebral columns with absent or rudimentary
intervertebral discs (Smits and Lefebvre, 2003; Choi and Harfe, 2011). Thus,
maintenance of cell separation between notochord and non-notochord derived cells is
integral to normal intervertebral disc morphogenesis and vertebral column formation.
At 14.5 dpc, Robo1 and Robo2 expression were observed in the presumptive
annuli fibrosi (Fig. 2-2). At a similar stage, Slit2 mRNA was found to be enriched in the
nuclei pulposi (Fig. 2-3). Based on these complementary expression patterns, it was
58
hypothesized that Robo-Slit signaling played a role in maintaining cell boundaries during
intervertebral disc development. Specifically, it was hypothesized that Slit secretion from
the nucleus pulposus repelled Robo-expressing annulus fibrosus cells from aberrantly
migrating into the nucleus pulposus (Fig. 2-8). Thus, in the absence of Robo1 and
Robo2 it was predicted that intervertebral discs in P0 mice would show signs of aberrant
cell mixing in the nuclei pulposi.
To determine if Robo1 and Robo2 in the presumptive annuli fibrosi played a role
in preventing migration of annulus fibrosus cells into the nucleus pulposus, ShhCre and
Rosa-mT/mG alleles were crossed into Robo1-/+;Robo2+/- mice (Fig. 2-6). ShhCre was
used to GFP-mark the notochord and all cells derived from the notochord. As in control
littermates, all cells in the nuclei pulposi of P0 Robo1;Robo2 null mice were GFP-
positive, indicating notochordal origin. Absence of tdTomato-positive cells in the nuclei
pulposi indicated that annulus fibrosus cells had not aberrantly migrated into the nuclei
pulposi during development. Additionally, as in controls, the vertebrae and annuli fibrosi
in Robo1;Robo2 null mice lacked GFP-positive cells, demonstrating that the notochord
had properly transitioned into nuclei pulposi anlagen during development (Fig. 2-6).
Together, these data show that the intervertebral disc defect seen in Robo1;Robo2 null
mutants was not caused by aberrant cell mixing in the nuclei pulposi as a result of
Robo1 and Robo2 removal.
Robo1;Robo2 null annuli fibrosi had aberrantly shaped cells that lacked the
distinct lamellar organization seen in control mice (Fig. 2-4). These changes were
accompanied by changes in the expression and distribution of extracellular matrix
proteins including Col2a1, Col6a1, and Aggrecan (Fig. 2-13). Col2a1, Col6a1, and
59
Aggrecan are important for cartilage function and reduced expression of Col2a1,
Col6a1, and Aggrecan are linked to intervertebral disc degeneration in humans and
animals (Smith, et al, 2011; Urban and Roberts, 2003; Sivan, et al, 2014).
Notably, one matrix protein that was found to be upregulated in Robo1;Robo2
null intervertebral discs was Tenascin C (Fig. 2-14) Tenascin C upregulation in the
intervertebral discs was accompanied by abnormal cell rounding in the annuli fibrosi of
Robo1;Robo2 null mice (Fig. 2-12). Tenascin C is a matricellular adhesion modulatory
protein that has been shown to impede cell-matrix adhesion by interfering with cell
interactions with fibronectin (Chiquet-Ehrismann, et al, 1988). Tenascin C has been
shown to cause cell rounding in in vitro studies (Huang, et al, 2001). Western blot
analyses of control and mutant littermate tissues showed that removal of Robo1 and
Robo2 resulted in an increase in Tenascin C protein expression in the discs (Fig. 2-14).
In these experiments, biological triplicates from control and mutant littermates were
used to quantify 170kD and 250kD Tenascin C fragments in the discs. Upregulation in
170kD and 250kD Tenascin C fragments has previously been linked to joint pathology
in humans (Sofat, et al, 2012). Additionally, recombinant 170kD and 250kD Tenascin C
fragments were previously shown to induce cartilage matrix degradation (Sofat, et al,
2012). Based on these results, the observation that Robo1;Robo2 null mice had
cartilage matrix protein deficiencies and aberrant cell rounding in the annulus fibrosus is
consistent with the observation that Tenascin C was upregulated in Robo1;Robo2 null
mutants.
60
Robo1 and Robo2 Function in the Sclerotome
Robo genes have been reported to regulate cell proliferation in vivo (Borrell, et al,
2012). Based on Robo1 and Robo2 expression in the sclerotome and the observation
that sclerotome-derived tissues (ribs) were hypoplastic at P0 (Fig. 2-8), 13.5 dpc
embryos were EdU pulsed for one hour and then examined for changes in cell
proliferation. Based on these experiments, removal of Robo1 and Robo2 was found to
impair proliferation in the sclerotome at 13.5 dpc (Fig. 2-9).
The proximal and distal ribs form from the sclerotome (Aoyama, et al, 2005).
Robo1;Robo2 null mutants had deficits in the ribs. Specifically, the distal ribs failed to
meet at the midline where the ribs fuse to the sternum (Fig. 2-8). The sternum derives
from the lateral plate mesoderm, which has been shown to require interactions with the
sclerotome for proper differentiation (Sudo, et al, 2001). In addition to deficits in the
distal ribs, Robo1;Robo2 null mutants sternal defects which included incomplete sternal
bar fusion and decreased sternal length (Fig. 2-8). It is not clear from these results if the
sternal defects were indirectly caused by Robo1 and Robo2 loss in the sclerotome, or if
Robo1 and Robo2 are expressed in and required for lateral plate mesoderm
differentiation.
Robo1 and Robo2 in Growth Plate Maintenance
Robo gene expression in the growth plate has been reported previously (Noel, et
al). Mechanical loading was found to increase Robo mRNA expression in the growth
plates, suggesting a role for Robo receptors in regulating growth plate chondrocyte
differentiation or homeostasis (Noel, et al, 1998). Xgal staining of vertebral columns
from Robo1+/-;Robo2+/- mice showed that Robo1 and Robo2 were expressed in the
61
vertebral growth plates at P0 (Fig. 2-11). To determine if Robo1 and Robo2 played a
role in regulating chondrocyte differentiation or function in the growth plates, RNA in situ
hybridization was used to evaluate expression of chondrocyte differentiation markers in
the growth plates.
Compared to control littermates which had distinct bands of Ihh and Col10a
expression in the vertebral growth plates, Robo1;Robo2 null growth plates had
markedly reduced Ihh and Col10a expression (Fig. 2-11). In the growth plates, Ihh and
Col10a are expressed by pre-hypertrophic and hypertrophic chondrocytes, respectively
(de Crombrugghe, et al, 2000). These gene expression changes were accompanied by
chondrocyte disorganization in the growth plates of Robo1;Robo2 null mice (Fig. 2-11).
While examining Ihh and Col10a expression in the growth plates, ectopic Ihh and
Col10a was observed in the annuli fibrosi of P0 Robo1;Robo2 null mice (Fig. 2-12).
Together, these data support a role for Robo receptors in regulating chondrocyte
maturation in the vertebral growth plates and suggest that the cell rounding phenotype
seen in Robo1;Robo2 null mutants may be caused by ectopic expression of
chondrocyte hypertrophy markers in the annuli fibrosi.
62
Figure 2-1. Robo1 and Robo2 Expression Analysis by RNA In Situ Hybridization and Xgal Staining. A) Robo expression analysis by RNA in situ hybridization and Xgal staining produces congruent results. Robo2 mRNA was detected in the neural tubes of wildtype 10.5 dpc embryos. B) Robo2 expression in the neural tube was confirmed by Xgal staining in stage matched Robo1;Robo2 heterozygotes. C) At 12.5 dpc, Robo2 mRNA appeared as repeating units in the somitic mesoderm along the rostrocaudal axis. D) Expression analysis by Xgal staining confirmed Robo2 expression in the vertebral column at 12.5 dpc. Wildtype embryos were used for RNA in situ hybridizations and Robo1;Robo2 heterozygotes were used for Xgal staining experiments. N=notochord; FP=floor plate
63
Figure 2-2. Robo Gene Expression During Vertebral Column Formation. A) Robo1 and Robo2 expression was assessed by Xgal staining and RNA in situ hybridization. At 12.5 dpc, Robo1 and Robo2 expression was observed as repeating segments in the somitic mesoderm lateral to the notochord. B) At 12.5 dpc Robo1 and Robo2 expression was observed in the somitic mesenchyme ventral to the notochord (red arrow). C) In wildtype 13.5 dpc embryos, Robo1 mRNA was found in the vertebral column just lateral to the midline. D) At 13.5 dpc, Robo2 mRNA was expressed at the midline, dorsal and ventral to the notochord. E) As assayed by Xgal staining, Robo1 and Robo2 were expressed in the spinal cord and vertebral column at 14.5 dpc. F) At 14.5 dpc Robo1 and Robo2 were expressed in the presumptive annuli fibrosi but were absent in the nuclei pulposi anlagen. G) At 18.5 dpc, Robo2 mRNA was expressed in the spinal cord, and to a lesser degree, in the vertebral column. H) 100um sections through the vertebral column showed that Robo1 and Robo2 were absent in the discs and were enriched in the vertebral growth plates (red arrows). I) In P0 vertebral columns, Robo1 and Robo2 were expressed in the trabecular centers of the vertebrae (red arrow), in the perichondrium (white arrowheads), and in isolated annulus fibrosus cells (red arrows). SpC=Spinal cord; VC=Vertebral column; NP=Nucleus pulposus; AF=Annulus fibrosus; V=Vertebra. Notochord denoted by black arrowheads in A, B, and D.
64
Figure 2-3. Slit Gene Expression During Vertebral Column Formation. A) RNA in situ hybridization using DIG-labeled antisense probes showed Slit1 gene expression in the notochord and floor plate at 10.5 dpc. B) At 10.5 dpc Slit2 mRNA was expressed in the floor plate and notochord. C) At 10.5 dpc Slit3 mRNA was observed in the floor plate. D) At 12.5 dpc Slit1 mRNA was expressed in the floor plate and notochord (black arrowhead). E) At 12.5 dpc Slit2 mRNA was expressed in the floor plate and in the notochord but was absent in the surrounding somitic mesoderm. F) At 12.5 dpc Slit3 was expressed in the floor plate but was not detected in the notochord. G) Slit1 mRNA was not detected in the intervertebral discs at 15.5 dpc. H) At 15.5 dpc Slit2 mRNA was enriched in the nuclei pulposi anlagen. I) At 15.5 dpc Slit3 mRNA was not observed in the intervertebral discs. FP=Floor plate; NP=Nucleus pulposus. Notochord denoted by black arrowheads in A-F.
65
Figure 2-4. Removal of Robo1 and Robo2 Causes Intervertebral Disc Malformations in P0 Mice. A) Removal of Robo1 and Robo2 resulted in enlarged, abnormally shaped discs. Compared to intervertebral discs from control littermates (a), mice null for Robo1 and Robo2 had enlarged, bulbous intervertebral discs (b). Disc malformations were apparent along the length of the thoracic and lumbar level regions (c, d). Robo1+/- and Robo1+/-;Robo2+/- controls were indistinguishable from wildtype littermates (not shown). B) Homozygous removal of Robo1 resulted in less severe disc malformations at P0. Robo1+/- control littermates had characteristically flattened discs (a) while Robo1 null mice had slight malformations in disc shape (b). C) Vertebral columns from 17.5 dpc Slit1-/-;Slit2-/- mice appeared comparable to those from control littermates. All images are of frontally sectioned vertebral columns that were stained with Alcian blue and Picrosirius red to show cartilage and fibrillar collagens, respectively. IVD=Intervertebral discs; NP=Nucleus pulposus.
66
Fig 2-5. Conditional Inactivation of Robo2F on Robo1 Null Background. A) X-gal stained Dermo1Cre;R26R tissues. At 9.5dpc Dermo1 was expressed in the notochord and surrounding mesoderm (a, b). At P0, Dermo1 showed mosaic expression in the vertebrae, annulus fibrosus, and nucleus pulposus (c). Embryos were sectioned across the transverse plane (a, b) or frontal plane (c). Tissues were Nuclear Red counterstained to show tissue architecture. B) Robo2F was inactivated in the somitic mesoderm using Dermo1Cre. Homozygous inactivation of Robo2F on a Robo1 null background partially recapitulated the Robo1;Robo2 null phenotype at P0. Vertebral columns from mutant and control animals were frontally sectioned and stained with Alcian blue and Picrosirius Red. Compared to controls (a), homozygous inactivation of Robo2F with Dermo1Cre on a Robo1 null background resulted in reduced Alcian blue staining in the vertebrae and intervertebral discs (b), signifying deficits in chondrogenesis following vertebral column formation. C) Robo2F inactivation with ShhCre on a Robo1 null background did not recapitulate the Robo1;Robo2 null phenotype. ShhCre is expressed in the notochord and is not expressed in the somitic mesoderm. NT=Neural tube; LB=Limb bud; FP=Floor plate; N=Notochord; NP=Nucleus pulposus; AF=Annulus fibrosus; V=Vertebrae.
67
Figure 2-6. The Notochord to Nucleus Pulposus Transition Occurs Normally in the Absence of Robo1 and Robo2. A) Model for Robo-Slit interaction during intervertebral disc morphogenesis. Slit and Robo exhibited complementary expression patterns in the vertebral column at 15.5 dpc with Robo1 and Robo2 expression in the presumptive annuli fibrosi (blue region) and Slit2 mRNA expression in the nuclei pulposi anlagen (red region) (a). Based on this model, in wildtype embryos Slit glycoprotein secretion from the nucleus pulposus would inhibit annulus fibrosus cells from migrating into nuclei pulposi anlagen via Slit protein interactions with Robo1 and Robo2 receptors expressed on annulus fibrosus cells (b). Removal of Robo1 and Robo2 would result in loss of Slit-mediated repulsion leading to a mixed population of nucleus pulposus and annulus fibrosus cells at birth. NP=Nucleus pulposus; AF=Annulus fibrosus. B) Mating scheme for lineage tracing analysis. The Rosa-mT/mG dual fluorescence reporter was used to assess the lineage of disc cells in perinatal vertebral columns. In the absence of Cre protein, all cells in the Rosa-mT/mG mouse fluoresce red (tdTomato). In the presence of Cre, Cre-mediated excision of the tdTomato cassette results in the production of membrane-targeted GFP protein. ShhCre was used to mark all cells in the embryonic notochord. All cells that derive from the notochord fluoresce green (GFP) in the perinatal vertebral column. Robo1+/-;Robo2+/-; ShhCre males were mated to Robo1+/-;Robo2+/-; Rosa-mT/mG females. Embryos were collected at 18.5 dpc and mice that were positive for both ShhCre and Rosa-mT/mG were selected for lineage analysis. C) Representative images from controls (a-h) and Robo1;Robo2 null mutants (i-p) showing localization of GFP+ cells within the nuclei pulposi (b, j) and exclusion of tdTomato+ non-notochord derived cells from the nucleus pulposus (a, i). Higher magnification of the nucleus pulposus – annulus fibrosus junction from a-d and i-l are shown in e-h and m-p, respectively. Scale bars = 100um (a-d, i-l); Scale bars = 50um (e-h, m-p). VB=vertebral body; AF=annulus fibrosus; NP=nucleus pulposus.
68
Figure 2-7. Lineage Tracing Analysis of ShhCreERT2 Marked Cells. A) Mating scheme for lineage tracing analysis, using Tamoxifen-inducible ShhCreERT2. Robo1+/-; Robo2+/-; Rosa-mT/mG females were mated to Robo1+/-; Robo2+/-; ShhCreERT2 males. Pregnant dams were administered a pulse of Tamoxifen by oral gavage on 10.5 dpc, and embryos were harvested for lineage analysis at 18.5 dpc. Using this scheme, all cells expressing Shh between 10.5dpc and approximately 12.0 dpc undergo Cre-mediated excision of the tdTomato cassette. B) Lineage tracing analyses by fluorescent microscopy. Derivatives of Shh-expressing cells reside in the nucleus pulposus at 18.5 dpc in controls (b, f) and Robo1;Robo2 null mutants (j, n). No ectopic tdTomato+ cells were found in the nuclei pulposi or either controls (a, e) or Robo1;Robo2 null mice (i, m). Higher magnification of the nucleus pulposus – annulus fibrosus junction shown in (e-h) and (m-p). Scale bars = 100um (a-h); Scale bars = 50um in (a‟-h)‟. VB=vertebral body; AF=annulus fibrosus; NP=nucleus pulposus.
69
Figure 2-8. Robo1;Robo2 Null Mice Have Deficits in the Distal Ribs and Sterna at P0. A) Robo1;Robo2 null mice have smaller rib cages at birth. Compared to controls (a-c), loss of Robo1 and Robo2 results in smaller ribs with underdeveloped distal ribs that are unfused to the sternum (d-g). All skeletal images were imaged at the same magnification (8x). B) Robo1 and Robo2 loss caused sternal defects. Representative skeletal preparations from one control (a) and two Robo1;2 null mutants (c, e) depict sternal fusion deficits caused by Robo1 and Robo2 removal. Higher magnification of the sterna in a, c, and e are shown in b, d, and f, respectively. Compared to control mice, which had distal ribs that attached at fused, segmented sterna (a, b), Robo1;Robo2 null mice had shorter sterna (c) that were unfused at the midline (d) and distal ribs that failed to stain with Alcian blue (e, f). C) Robo1;2 null mutants had a 30% decrease in sternal length at P0 compared to controls. n=2 controls; n=5 Robo1;Robo2 null mutants.
70
Figure 2-9. Impaired Proliferation in Robo1; Robo2 Null Mutants at 13.5 dpc. A) Anti-β-Galactosidase (bGal) antibody is a reliable proxy for Robo expression in the vertebral column. Sagittal cross sections from a 14.5 dpc Robo1+/-;Robo2+/- embryo shows Robo expression by Xgal staining in the vertebral column (a). Sagittal sections from 13.5 dpc Robo1+/-;Robo2+/- mice were incubated with anti-bGal primary antibody and 555-conjugated secondary antibody to show bGal protein in the vertebral column (b). Both images are shown at 20x. B) Schematic illustrating how percent proliferation in the sclerotome was quantified. Sagittal cross sections through the midline from the trunk level of 13.5 dpc embryos were used to fluorescently label nuclei (c). An antibody against β-galactosidase was used as a proxy for Robo1/2 expression in the vertebral column (b). Percent proliferation was calculated as the total number of EdU positive
71
cells within β-Gal-488 positive domains (d) divided by total number of DAPI nuclei within the same β-Gal-488 domains (c). SpC = Spinal cord; Noto = Notochord. C) Representative images from control (a-d) and Robo1;2 mutants (e-h) showing proliferation in trunk region at 13.5 dpc. Vertebral columns were sectioned sagittally and are shown in the same orientation, with spinal cord on the left. Midline sections in the trunk region between the fore- and hindlimbs were selected for analysis. Scalebar = 100um. D) Compared to controls, which showed 23.6 % proliferation in the Robo-expressing sclerotome, Robo1/2 null embryos showed 11.5% proliferation in the sclerotome. Percent proliferation in the notochord, which does not express Robo, was used as a control for overall mitotic index. No change in proliferation in the notochord was observed. Pregnant dams were EdU pulsed for one hour by IP injection. Robo1+/-;Robo2+/- littermates were used as controls to evaluate the effect of Robo loss on proliferation. n=3 controls, n=3 mutants.
72
Figure 2-10. Robo and Pax1 Have Overlapping Expression Patterns in the Developing Vertebral Column. A) Pax1 and Robo1;Robo2 have similar but not identical expression patterns in the developing vertebral column. Robo1+/-;Robo2+/- embryos were Xgal stained to show Robo1 and Robo2 in the vertebral column at 12.5 dpc and 14.5 dpc (a, e). An antisense probe to Robo2 mRNA was used to show Robo2 expression in wildtype 13.5 dpc embryos (c). Anti-Pax1 antibody was used to show Pax1 expression in stage-matched wildtype or Robo1+/-;Robo2+/- embryos (b, d, f). SpC=spinal cord. Merged Pax1 and DAPI immunofluorescence shown in b, d, f. B) Pax1 and Robo are co-expressed in a small subset of cells in the sclerotome. Sagittally sectioned tissue from a 13.5 dpc Robo1+/-; Robo2+/- embryo was fluorescently labeled for Pax1 (a) and bGal (b). bGal was used as a proxy for Robo expression. Z-stacked images taken by confocal microscopy show that Pax1 and Robo have similar expression patterns in the vertebral column (a, b), and that a subset of cells in the ventral sclerotome co-express Pax1 and Robo at this stage (c).
73
Figure 2-11. Disrupted Growth Plate Marker Expression in Robo1;Robo2 Null Mutants. A) Chondrocytes in the vertebral body (VB) growth plates appear disorganized in Robo1;Robo2 null mutants at P0 (d) compared to control littermate growth plates, which have distinct layers of small, quiescent cells and larger, more differentiated cells nearer to the VB ossification centers (b). RZ=Resting zone; PZ=Proliferative zone;
74
HZ=Hypertrophic zone; TB=Trabecular bone. All images are of frontally sectioned, Alcian blue and Picrosirius red stained vertebral columns from P0 mice. (B) In perinatal vertebral columns, Robo1;Robo2 expression, as assayed by Xgal, is enriched in the VB growth plates. C) Expression of chondrocyte differentiation markers is disrupted in the vertebral growth plates of Robo1;Robo2 null mice at P0. Ihh expression appeared reduced in Robo1;Robo2 null mutants at P0 (b) compared to control littermates (a) which had Ihh mRNA enriched at the growth plates. Col10a, which is expressed by hypertrophic chondrocytes, was barely detectable in Robo1;Robo2 null vertebral columns (d). By comparison Col10a expression was specific to and enriched in the vertebral growth plates (c). VB=Vertebral body; NP=Nucleus pulposus. Growth plates denoted by white arrows.
Figure 2-12. Robo1;Robo2 Null Annuli Fibrosi Have Aberrant Cell Morphologies And Ectopic Expression of Growth Plate Chondrocyte Markers. A) Robo1;Robo2 null
75
mutants show aberrant cell morphology in the annuli fibrosi at P0. Compared to annulus fibrosus cells in control mice which are spindle shaped and organized into concentric rings around the nucleus pulposus (a), annulus fibrosus cells in Robo1;Robo2 null mice appear rounded and enlarged (b). B) These cell shape changes are accompanied by ectopic expression of chondrocyte hypertrophy and pre-hypertrophy markers in the discs. While Ihh expression is confined to the vertebral growth plates in controls (a), Ihh is ectopically expressed by annulus fibrosus cells in Robo1;Robo2 null mice (b, red arrow). Col10a1 expression is restricted to hypertrophic growth plate chondrocytes in controls (c). Robo1;Robo2 null mice show loss of an organized hypertrophic zone and express Col10a1 ectopically in the annulus fibrosus (d, red arrow). All images (A, B) are frontal cross-sections from P0 mice.
Figure 2-13. Robo1 and Robo2 Loss is Associated With Reduced Expression of Cartilage Matrix Proteins. A) Col6a1 is decreased in the intervertebral discs of Robo1;Robo2 null mice. Compared to discs from control littermates (a), discs from Robo1;Robo2 null mice show decreased staining for Col6a1 (b). B) Col2a1 expression is enriched in the inner annuli fibrosi. At P0, Col2a1 immunofluorescence in control and Robo1;Robo2 null littermates appears comparable. In control (a) and Robo1;Robo2 mutant discs (b) Col2a1 is enriched in the inner annuli fibrosi. C) Aggrecan is decreased in Robo1;Robo2 null mutants. Compared to discs from control littermates which had robust staining throughout the nucleus pulposus and annulus fibrosus (a), Robo1;Robo2 null discs had reduced staining in the annulus fibrosus (b). All images are of frontally sectioned vertebral columns from P0 mice and are shown at 20x. af=annulus fibrosus; np=nucleus pulposus. DAPI shown in c, d; merged images shown in e, f.
76
Figure 2-14. Tenascin C is Upregulated in the Intervertebral Discs of Robo1;Robo2 Null mice. A) Tenascin C upregulation in Robo1;Robo2 null mutant discs can be seen by immunofluorescence. Frontally sectioned discs from P0 control and Robo1;Robo2 null littermates were incubated with anti-Tenascin C antibody. In both control and Robo1;Robo2 null discs, Tenascin C is enriched in the annuli fibrosi (a, b). DAPI shown in c, d. Tenascin C/DAPI merged images shown in e, f. All images are 20x. B) Western blot analysis of Tenascin C in P0 discs. Thoracic and lumbar level discs from P0 mice were pooled for protein quantification by western blot. Each lane represents one biological replicate (10-14 discs pooled from the same animal). Compared to discs from wildtype littermates (lanes 1-3), discs from Robo1;Robo2 null mice showed an increase in high molecular weight Tenascin C (lanes 4-6). C) Western blot quantification. Robo1;Robo2 null mutants had a 60% increase in Tenascin C in the discs at P0. GAPDH was used as a loading control to calculate fold change based on densitometry of the 170kD and 250kD Tenascin C bands.
77
Figure 2-15. Slit1;Slit2 Null Phenotype. Deletion of Slit1 and Slit2 did not recapitulate the Robo1;Robo2 null intervertebral disc phenotype. Vertebral columns from 17.5 dpc Slit1+/-;Slit2+/- control littermates (A, C) and Slit1;Slit2 null mutants (B, D) were frontally sectioned and stained with Alcian blue and Picrosirius Red. Annulus fibrosus cell morphologies were normal in Slit1;Slit2 null mice at 17.5 dpc (C-D). NP=Nucleus pulposus; AF=Annulus fibrosus; V=Vertebra
78
CHAPTER 3 WNT/β-CATENIN SIGNALING DURING INTERVERTEBRAL DISC DEVELOPMENT
Introduction
The Wnt/β-Catenin signaling pathway is an evolutionarily conserved signal
transduction network that mediates many crucial developmental and disease processes,
including intervertebral disc formation, maintenance, and intervertebral disc disease as
well as cartilage and long bone development (Dahia, Mahoney, et al. 2009; Winkler,
Mahoney, et al. 2014; Usami, Gunawardena, et al. 2016). In the canonical Wnt/β-
Catenin signaling pathway, secreted Wnt glycoproteins bind to Frizzled (Fz)
transmembrane G protein-coupled receptor proteins, resulting in β-Catenin
accumulation in the cytoplasm followed by β-Catenin translocation into the nucleus. In
the nucleus, β-Catenin proteins interact with TCF/Lef family of transcription factors to
regulate expression of downstream targets (Cadigan and Nusse).
In the early embryo within the axial mesoderm, Wnt/β-Catenin signaling is
required for notochord formation and extension, and in mice inactivation of the Wnt/β-
Catenin signaling pathway is known to cause deficits in notochord elongation (Cheyette,
Waxman, et al. 2002; Ukita, et al. 2009). Following notochord formation, opposing
gradients of Shh and Wnt signaling in the somitic mesoderm confer dorsal-ventral
polarity to the somitic mesoderm for induction of the dermamyotome and sclerotome,
respectively (Fan, Lee, et al. 1997; Fan, Tessier-Lavigne, 1994). In vitro, pluripotent
cells can be induced to form sclerotome-like chondroprogenitors when treated with
drugs that modulate the Wnt/β-Catenin signaling pathway (Zhao, Li, et al. 2014).
Recent studies in vertebrate models implicate Wnt/β-Catenin signaling in
intervertebral disc development and disease. In 18.5dpc mice Wnt signaling has been
79
observed in and near the intervertebral discs, including the annulus fibrosus,
cartilaginous endplates, and vertebral growth plates (Kondo, et al. 2012). In these
studies, Wnt/β-Catenin signaling was not observed in the nuclei pulposi during
embryonic stages, but was found to be up-regulated in the nucleus pulposus
postnatally, concomitant with Wnt/β-Catenin down-regulation in the annulus fibrosus
and complete loss of Wnt/β-Catenin signaling in the growth plates and endplates
(Kondo, et al. 2012). Another study that used immunohistological methods similarly
found evidence of Wnt signaling activity in the intervertebral discs of prenatal and
postnatal mice (Dahia, Mahoney, et al. 2009).
Changes in Wnt/β-Catenin signaling activity have also been linked to
intervertebral disc degeneration in vertebrate animal models. In rats, chemically induced
nuclear accumulation of β-Catenin in the intervertebral discs has been shown to induce
cell senescence and apoptosis in the nucleus pulposus (Hiyama, Sakai, et al. 2010). A
decrease in canonical Wnt signaling and caveolin-1 expression has also been linked to
intervertebral disc disease in chondrodysplastic dogs (Smolders, et al. 2013). In mice,
Wnt/β-Catenin signaling activity was found to be reduced in the intervertebral discs of
aged animals and was linked to degenerative changes in the intervertebral discs,
including decreases in cell proliferation and reduced expression of chondrogenic
markers, including Sox9, Aggrecan, Col2a1, and Col1a1 (Winkler, Mahoney, et al;
Hiyama, Sakai, et al). Additionally, activation of the Wnt/β-Catenin signaling pathway
has been shown to promote intervertebral disc degeneration by inducing expression of
matrix degrading enzymes (Hiyama, Sakai, et al).
80
Previous studies that evaluated Wnt/β-Catenin signaling in the intervertebral
discs have produced variable and conflicting results. These results were produced using
immunohistological staining methods and Xgal staining of TOPGAL reporter mice. To
evaluate Wnt/β-Catenin signaling activity in the intervertebral discs using a novel
method, TCF/Lef-H2B-GFP reporter mice were used to characterize Wnt/β-Catenin
signaling activity in the vertebral column and intervertebral discs during embryonic and
postnatal development.
Results
TCF/Lef:H2B-GFP reporter mice (Ferrer-Vaquer, et al.) were used to investigate
Wnt/β-Catenin signaling activity in the developing vertebral column. In the canonical
Wnt signaling pathway, Wnt activation of the Fz G protein-coupled cell surface receptor
results in stabilization of cytoplasmic β-Catenin and translocation of β-Catenin to the
nucleus where it interacts with T cell-specific transcription factor/lymphoid enhancer-
binding factor 1 (TCF/Lef) family of transcription factors to drive expression of Wnt-
responsive genes (MacDonald, et al. 2009). The TCF/Lef:H2B-GFP reporter used in
these experiments were generated using a construct consisting of H2B-GFP under the
control of six TCF/Lef response elements and the hsp68 minimal promoter. Nuclear
localization of the chromatin-bound H2B-GFP fusion protein was designed to enable
GFP readout at a single-cell resolution (Ferrer-Vaquer, et al. 2010).
At 12.5 dpc, Wnt/β-Catenin signaling activity was observed in the tail-level
notochord in a region caudal to the external genitalia (Fig. 3-1B). Wnt signaling was also
observed in the more anterior trunk-level notochord, between the fore- and hindlimbs
(Fig. 3-1E). These data were consistent with published results that reported Wnt
81
signaling activity in the notochord (Maretto, Cordenonsi, et al. 2003) and with reports
that demonstrated a role for Wnt/β-Catenin signaling in notochord extension and tail
formation (Ukita, Hirahara, et al. 2009).
Following the notochord to nucleus pulposus transition at 15.5 dpc, Wnt/β-
Catenin signaling activity was detected in the subset of nucleus pulposus cells in the
vertebral column (Fig. 3-2B and Fig. 3-2D). Wnt/β-Catenin signaling was not observed
in annulus fibrosus cells at this stage (Fig. 3-2A-D). At 18.5dpc, Wnt signaling was
maintained in the spinal cord, as reported previously (Ferrer-Vaquer, et al. 2010) and in
a small number of isolated chondrocytes located in the vertebrae (Fig. 3-2F). Wnt
signaling was also observed in isolated nucleus pulposus cells at 18.5 dpc (Fig. 3-2F
and Fig. 3-2H). These data are in contrast to published studies that showed absence of
Wnt/β-Catenin signaling activity in the nucleus pulposus at 18.5dpc (Kondo, et al). At
18.5dpc Wnt/β-Catenin signaling activity was reported in the annulus fibrosus and
endplates (Kondo, et al, 2011).
In postnatal mice, Wnt/β-Catenin signaling was maintained in the spinal cord
(Fig. 3-3C, Fig. 3-3L). In one-month-old postnatal mice, Wnt/β-Catenin signaling was
detected in a subset of nucleus pulposus cells (Fig. 3-3E). These data are in agreement
with a previously published study that reported Wnt/β-Catenin signaling in the nucleus
pulposus at five weeks old (Kondo, et al, 2011). At one month old, very little Wnt
signaling activity was observed in the annulus fibrosus (representative images of the
annulus fibrosus at one month shown in Fig. 3-3G-I). A few isolated GFP-positive cells
were detected in the annulus fibrosus at one month old (not shown) and in perichondrial
mesenchymal cells located at the periphery of the vertebral column (Fig. 3-3H). These
82
data are also in agreement with published data that showed Wnt signaling down-
regulation in the annulus fibrosus in comparably aged postnatal mice (Kondo, et al.
2011). Finally, at one year old, Wnt signaling was markedly decreased in all parts of the
intervertebral discs but was observed in trabecular osteoblasts located in the vertebral
bodies (Fig. 3-3J-L).
Conclusions
Wnt/β-Catenin signaling has been implicated in intervertebral disc development
in disease in humans and animals. The status of Wnt/β-Catenin signaling in
intervertebral disc and disc progenitor cells have been investigated using a variety of
methods, including evaluation of β-Catenin mRNA and protein and LacZ-based reporter
methods (Hiyama, et al. 2010; Dahia, et al. 2009; Winkler, et al. 2014). These studies
have often produced inconsistent results. To assess the status of Wnt signaling using a
novel method, TCF/Lef-H2B:GFP reporter mice were used to monitor Wnt/β-Catenin
signaling in the notochord and intervertebral discs at a single-cell resolution at
embryonic and postnatal stages.
Robust GFP fluorescence, indicative of high levels of Wnt signaling activity, was
detected throughout the notochord at 12.5 dpc, congruent with published results that
demonstrated a role for Wnt/β-Catenin signaling in notochord (Ukita, Hirahara, et al.). At
15.5 dpc, after the notochord had transitioned into nuclei pulposi, Wnt/β-Catenin activity
was detected in approximately 50% of cells located in the nuclei pulposi. These data
were congruent with a published study that showed nuclear β-Catenin protein staining in
the nucleus pulposus of 15.0dpc mouse embryos (Hiyama, Sakai, et al).
83
In 18.5 dpc and in one month old postnatal mice, Wnt signaling was observed in
the nuclei pulposi, albeit at lower levels than at 15.5 dpc (Fig. 3-2 and Fig. 3-3).
Compared to intervertebral discs at 18.5 dpc, tissues from one month old postnatal mice
showed GFP fluorescence in the nuclei pulposi, albeit in a smaller percentage of cells
(Fig. 3-3). These data are in contrast to a published report that showed upregulation in
Wnt/β-Catenin signaling in the nuclei pulposi between 18.5 dpc and five weeks old
(Kondo, et al. 2011). The reporter used in those studies was the TOPGAL reporter
strain, in which LacZ transgene expression is driven by three TCF/LEF binding motifs
upstream of a minimal c-fos promoter (DasGupta and Fuchs, 1999). Another study,
which also used the TOPGAL Wnt reporter, reported that in postnatal intervertebral
discs (one week and three weeks old) Wnt signaling activity was present in the annuli
fibrosi but not in the nuclei pulposi (Dahia et al. 2009). These results are inconsistent
with the observations made using the TCF/Lef:H2B-GFP reporter.
Postnatal intervertebral discs from one month old mice showed Wnt/β-Catenin
signaling in the nucleus pulposus. Intervertebral discs from one year old mice showed
markedly reduced or absent Wnt/β-Catenin signaling in the nuclei pulposi and annuli
fibrosi (Fig. 3-3). Thus, Wnt/β-Catenin signaling is down-regulated in the intervertebral
discs with age. These results are congruent with published data that suggested that
Wnt/β-Catenin signaling is down-regulated in the intervertebral discs as animals age
(Winkler, et al. 2014). Down-regulation in the Wnt/β-Catenin signaling pathway may be
important for postnatal intervertebral disc homeostasis. Mice that had conditional re-
activation of the Wnt/β-Catenin signaling pathway showed signs of degenerative
changes in the discs (Wang, et al. 2013) and postnatal activation of the Wnt/β-Catenin
84
signaling pathway was shown to enhance intervertebral disc cell senescence in rats
(Hiyama, et al. 2013).
In sum, these data confirmed the presence of Wnt/β-Catenin signaling in the
embryonic notochord, where it has been shown to regulate notochord extension (Ukita,
et al 2009). These data also suggest that Wnt signaling may be more integral to nucleus
pulposus development than annulus fibrosus development, and show that Wnt signaling
is down-regulated in the discs with age. Dissimilarities between our data and published
results may be explained by the use of different methods to assess Wnt signaling
activity. Previous studies relied on the assessment of β-Catenin protein or mRNA
localization or on the use of TOPGAL reporter mice, which relied on LacZ driven by
three TCF/Lef binding motifs and c-fos promoter.
Collectively these data highlight the utility of the TCF/Lef-H2B-GFP mouse for
reporting Wnt signaling activity in postnatal intervertebral discs, when Wnt/β-Catenin
signaling activity may be too low to detect by other methods. These data support a role
for Wnt/β-Catenin signaling in intervertebral disc development, and suggest that Wnt/β-
Catenin signaling down-regulation at postnatal stages may be a normal facet of disc
biology and homeostasis.
85
Figure 3-1. WNT/β-Catenin Signaling in the Notochord at 12.5 dpc. Wnt signaling was observed in the notochord at 12.5 dpc in the caudal notochord located in the tail (A-C white arrows) and in the trunk-level notochord between the fore- and hindlimb buds (D-F white arrows). Wnt signaling was also observed in neuronal progenitors in the neural tube (NT). Embryos were sectioned through the sagittal plane, counterstained with DAPI, and imaged at 20x. NT=Neural tube; N=Notochord; EG=External genitalia.
86
Figure 3-2. Wnt Signaling in the Embryonic Intervertebral Discs. WNT/β-Catenin Signaling was observed in the intervertebral discs at 15.5 dpc (A-D white arrows) and 18.5 dpc (E-H white arrows). At 15.5 dpc, WNT/β-Catenin signaling was observed in the neural tube and in the nuclei pulposi of the intervertebral discs (B, white arrows). Higher magnification of the panel C inset shows WNT signaling in a subset of nucleus pulposus cells, which are circumscribed in white dotted lines. No Wnt signaling was observed in the presumptive annulus fibrosus at 15.5 dpc. At 18.5 dpc, WNT signaling was observed in the vertebral column (E-H white and red arrows). Specifically, WNT signaling was detected in intervertebral disc cells (F, white arrow) and in chondrocytes located in the vertebrae (F, red arrow). A higher magnification GFP/DAPI image shows that WNT signaling was specific to nucleus pulposus cells (H, white arrows). No WNT signaling was observed in the annulus fibrosus at 18.5 dpc. The nucleus pulposus-annulus fibrosus boundary is denoted by the dashed white line in H. Vertebral columns were sectioned through the sagittal plane, counterstained with DAPI, and imaged at 20x (A-C, E-G).
87
Figure 3-3. Postnatal Wnt Signaling in the Intervertebral Discs. Sagittal cross sections from one month old postnatal TCF/Lef:H2B-GFP mice showed Wnt signaling in the nuclei pulposi of the intervertebral discs, denoted by white dashed lines in A-C. The nucleus pulposus and annulus fibrosus are denoted by the white and red boxes,
88
respectively, in B-C. Higher magnification images of the nucleus pulposus, denoted by the white box, are shown in D-F. The annulus fibrosus, denoted by the red box, are shown at higher magnification in G-I. Wnt signaling was observed in isolated cells located in the nucleus pulposus (E, white arrows). Very little Wnt signaling was observed in the annulus fibrosus at one month (G-I). GFP-positive cells were observed in perichondrial mesenchyme in one month old mice (H, red arrows). At one year postnatal, Wnt signaling was undetectable in the intervertebral discs but was observed in vertebral osteoblasts (J-L, red arrows). SpC=Spinal cord; V=Vertebrae; GP=Growth plate; NP=Nucleus pulposus; AF=Annulus fibrosus. Scalebar=100um (A-F, J-L) and 50um (G-I).
89
CHAPTER 4 METHODS
Histology
Vertebral column tissues were stained with Alcian blue and Picrosirius red as
described previously1, 2 (Harfe, Choi PNAS1; Gruber, et al 2002). Briefly, 4% PFA-fixed
tissues were washed in PBS and dehydrated through a series of graded ethanol
washes (water, 25% EtOH, 50% EtOH, 70% EtOH, 95% EtOH, 100% EtOH) before
being immersed in xylenes in preparation for paraffin embedding. Tissues were
perfused with Blue Ribbon paraffin and embedded in paraffin at 65°. A microtome was
used to section vertebral columns at 10um onto SuperFrost Plus Gold glass slides.
Slides were left to dry 24-72 hours on a slide warmer.
Tissues were re-hydrated through xylenes and ethanol washes and then
immersed in Alcian blue pH 2.5 for 15 minutes. Following Alcian blue, slides were
washed in running tap water to prepare for Picrosirius red. Slides were immersed in
Picrosirius red for 45 minutes, washed in acidified water (0.025% acetic acid). Slides
were then put through a series of ethanol washes again in preparation for mounting with
Permount.
Skeletal Preparations
Alcian blue and alizarin red were used to stain cartilage and ossified skeletal
elements as described previously3. Embryos were skinned, eviscerated, and fixed
overnight in 4% PFA/PBS. After fixation, tissues were PBS washed and dehydrated
through a series of ethanol washes in preparation for Alcian blue. Skeletons were
submersed in Alcian blue overnight at room temperature. The next day excess Alcian
blue was removed with multiple washes through 30% acetic acid/70%ethanol. Tissues
90
were then rehydrated and stained overnight in alizarin red in 1% KOH. Following alizarin
red, skeletons were cleared for several days in 1% KOH and then stored and imaged in
80% glycerol.
Western Blot Analysis
Vertebral columns were dissected into cold PBS, and then thoracic and lumbar
level discs were microdissected out and pooled into 200 uL of RIPA
(radioimmunoprecipitation assay buffer) lysis buffer supplemented with PMSF (final
concentration of 1mM) and EDTA (0.5M) for each animal. Discs were homogenized with
an electric homogenizer on ice. Samples were centrifuged at 13,000rpm for 12 minutes
and supernatant was placed in a clean tube. BCA assay (BioRad) was used to
determine protein concentration and all samples were brought to equimolar
concentration with RIPA buffer. Samples were prepared with 6x Laemmli loading buffer
and heated to 95° for five minutes.
20ug total lysate from each sample was loaded into 4-20% Mini-PROTEAN TGX
Precast Protein Gels purchased from BioRad and standard running buffer (1x Tris
glycine) was used for gel electrophoresis. Transfer buffer (20% final concentration
MeOH) was used for wet transfer proteins onto PVDF membrane at 100V for two hours
at 4°. Blots were blocked with 3% BSA for one hour at room temperature before
incubation with primary antibody. Tenascin C blots were incubated with 1:1000 anti-
Tenascin C antibody from Santa Cruz (H-300), diluted in 3% BSA, overnight at 4°.
PBST (0.1% Tween) was used to wash blots 3x at room temperature. Blots were then
incubated at room temperature in secondary antibody, which was Anti-Rabbit-HRP
secondary antibody diluted 1:1000 in 3% BSA. Blots were left in secondary antibody at
91
room temperature for one hour, or overnight at 4°. After Tenascin C detection, blots
were stripped with mild stripping buffer and GAPDH (1:2000) was applied next. GAPDH
(6C5) antibody was purchased from Abcam.
Protein bands were detected using ECL chemiluminescence on film, and ImageJ
software was used to determine, based on densitometry, changes in Tenascin C
relative to GAPDH bands. Each lane represents one biological replicate, n=3 for each
genotype. Each biological replicate is defined as 10-14 pooled discs from the thoracic
and lumbar regions.
ECM Analysis By Immunofluorescence
The relative quantity and localization of matrix proteins was examined by
immunofluorescence using antibodies against proteins that are enriched in cartilaginous
tissues9-11. Vertebral columns were dissected in cold PBS and fixed overnight in 4%
PFA/PBS at 4°. The next day, tissues were washed in PBS and cryopreserved by
immersing in 30% sucrose/PBS overnight at 4°. Tissues were embedded in Tissue-Tek
OCT (Optimal Cutting Temperature) embedding compound on dry ice and then stored
at -80°. A cryostat was used to section embryos and vertebral columns onto Superfrost
Plus Gold Slides. All sections were 20um unless otherwise specified.
Slides were retrieved from -80° and allowed to thaw to room temperature. A
PAP-Pen was used to draw a hydrophobic barrier around the tissues. Tissues were
permeabilized with three washes of PBST and then blocked for one hour at room
temperature in 3% BSA dissolved in PBST. Unless otherwise specified, all primary
antibodies were diluted 1:200 in 3% BSA/PBST. Tissues were incubated with primary
antibody overnight at 4° in a humidified slide box. The next day, tissues were washed
92
three times with PBST and then incubated with secondary antibody at room
temperature for one hour, or overnight at 4°, protected from light. Secondary antibodies
were diluted 1:200 in 3% BSA/PBST. After secondary antibody incubation, slides were
counterstained with DAPI nuclear stain. After a final round of PBST washes, slides were
coverslipped using Dako fluorescent mounting media and #1.5 glass coverslips. All
slides were stored at 4°, protected from light, and imaged on a Leica confocal
microscope. Images presented in figures are representative of multiple discs from the
thoracic and lumbar regions from at least two controls and two Robo1;2 null mice.
Characterization Of Robo And Slit Expression
X-Gal Staining
LacZ knockin into the Robo1 and Robo2 loci was used to generate the Robo1
and Robo2 null alleles. This allowed us to use Xgal staining to show Robo expression in
the vertebral column. Importantly, because both Robo1 and Robo2 were replaced by
LacZ, X-gal staining is a proxy for Robo1 and/or Robo2 expression. Vertebral columns
from Robo1+/-;Robo2+/- mice were indistinguishable from vertebral columns from
wildtype (Robo1+/+;Robo2+/+) mice. Thus, Robo1+/-:Robo2+/- embryos were used for
Xgal staining experiments. Staining was done as previously described4 (Harfe et al.
2004).
Skinned embryos (14.5 dpc and younger) were fixed overnight in 0.2% PFA.
Mice older than 14.5 dpc were additionally eviscerated before fixation. The next day,
embryos were prepared for X-gal staining with three washes in PBS. Tissues were
stained overnight at room temperature, protected from light. Where applicable, wildtype
embryos and tissues were used as negative controls for endogenous b-galactosidase
93
activity in the skeleton. After staining, tissues were washed in PBS and then fixed in 4%
PFA/PBS. X-gal substrate was prepared as 20mg/ml in DMF and used at a final
concentration of 1mg/ml.
Perinatal vertebral columns were additionally decalcified in Cal-Ex Decalcifier
overnight at room temperature and then washed in PBS. For sectioning, tissues were
either prepared for vibatome sectioning (100um) or microtome sectioning (10um).
Tissues that were vibratome sectioned were embedded in 6% agarose on ice after
overnight incubation in 30% sucrose. Tissues that were microtome sectioned were
prepared for paraffin embedding as described under Histology, above.
RNA In Situ Hybridization
All harvests and dissections for in situ hybridization experiments were done with
RNase Zap cleaned tools and work surfaces to minimize RNA degradation. Buffers
were DEPC-treated and kept on ice during dissections and incubations. Embryos were
harvested and dissected in cold DEPC-PBS and fixed overnight in 4% PFA prepared in
DEPC-PBS. Tissues were cryopreserved in 30% sucrose prepared in DEPC-PBS and
embedded on dry ice using OCT embedding media. Unless otherwise specified, tissues
were sectioned on a cryostat at 20um onto Superfrost Plus Gold glass slides and stored
at -80°. Following in situ hybridization, tissues were fixed in 4% PFA, PBS washed, and
mounted with Glycergel (Dako).
RNA probe templates for Robo1, Robo2, and Slit1-3 were generously provided
by Dr. Xin Sun5. DIG labeled probes were made from cDNA templates as previously
described6 with minor modifications. Briefly, DIG labeled RNA probes were synthesized
94
by in vitro transcription using DIG RNA Labeling kit (Roche). Probes were purified using
Mini Quick Spin RNA columns (Roche) and then stored at -80°.
RNA in situ hybridization experiments were conducted as previously described7
with minor modifications. Briefly, on day 1 tissues mounted on slides were retrieved
from -80° and allowed to come to room temperature. A PAP pen was used to
circumscribe tissues with a hydrophobic barrier before tissues were put through a series
DEPC-PBST washes in preparation for probe incubation. Tissues were postfixed in 4%
PFA for 20 minutes at room temperature and incubated with prehybe (10% dextran
sulfate) for one hour at 65°. Probes were diluted into 100ul fresh, pre-warmed prehybe
before being added to tissues for overnight incubation at 65°. Tissues were carefully
covered with parafilm and incubated in clean, humidified slide boxes to prevent
evaporation during overnight hybridization. On day 2, slides were put through a series
SSC/formamide washes at 65° to remove any excess, unhybridized probe. In
preparation for antibody incubation, slides were brought to room temperature in KTBT
and then blocked for one hour at room temperature in 20% HINGS (heat-inactivated
goat serum, 20% by volume in KTBT). After blocking, tissues were incubated at 4°
overnight with an AP-conjugated anti-DIG antibody (Roche) diluted 1:2000 in 20%
HINGS. On day 3, slides were washed 6x in KTBT at room temperature, and then left in
KTBT overnight at 4°. Hybridized tissues were developed with either BM Purple (Roche)
or with BCIP/NBT (final concentration 0.175mg/ml for BCIP and 0.225mg/ml for NBT).
Both reagents are substrates for alkaline phosphate, which produces a purple (BM
Purple) or blue (BCIP/NBT) precipitate that can be visualized by light microscopy. Slides
95
were PBS washed and postfixed in 4% PFA before being mounted with Glycergel
(Dako) for imaging.
Lineage Tracing Analyses
The Rosa-mT/mG dual fluorescent reporter8 allele was used to lineage trace the
origin of cells in the P0 vertebral column. ShhCre4 was used to mark notochord-derived
cells in the nucleus pulposus. For lineage tracing studies, P0 mice were collected from
Robo1+/-;Robo2+/-;Rosa-mT/mG females mated to Robo1+/-;Robo2+/-;ShhCre males
and prepared for cryostat sectioning. Briefly, vertebral columns were dissected into cold
PBS and fixed overnight in 4% PFA/PBS. Tissues were then washed in PBS and
submersed in 30% sucrose overnight. The next day, vertebral columns were embedded
on dry ice using OCT embedding media. Tissues were sectioned at 20um on a cryostat.
For imaging, tissues were counterstained with DAPI nuclear stain and mounted using
Dako fluorescent mounting media. Images were taken on a Leica confocal microscope.
For lineage tracing of notochord cells at 10.5dpc, the ShhCreERT24 allele was
used to selectively mark Shh expressing cells between 10.5dpc and 12dpc. In these
experiments, Robo1+/-;Robo2+/-;ShhCreERT2 males were mated to Robo1+/-
;Robo2+/-;Rosa-mT/mG females. The appearance of a vaginal plug was designated as
embryonic day 0.5 (0.5dpc). Tamoxifen dissolved in corn oil was administered by oral
gavage at 10.5dpc and then pups were collected at P0. Tissues were prepared for
analysis as described above.
96
Analysis Of Sclerotomal Proliferation
EdU Pulse And Detection
Time mated Robo1+/-;Robo2+/- females crossed to Robo1+/-;Robo2+/- males were
used to evaluate the effect of Robo loss on sclerotome proliferation. Analysis of EdU
proliferation in skeletal tissues was performed as described previously12. Briefly,
pregnant dams were weight at 13.5dpc for EdU injection by IP. Embryos were
harvested after one hour. For analysis of 18.5dpc tissues, EdU pulse was extended to
two hours.
Embryos and vertebral columns were dissected in PBS and fixed overnight in 4%
PFA/PBS at 4°. Robo1+/-;Robo2+/- control and Robo1-/-;Robo2-/- mutant littermates were
selected for further processing. Embryos were cryopreserved in 30% sucrose/PBS
overnight and then embedded in OCT embedding medium on dry ice. Sagittal sections
from 13.5 dpc embryos were sectioned at 20um on a cryostat, and sections through the
medial vertebral column containing notochord and/or discs were selected for further
analysis.
For detection of EdU incoporation, Click-iT EdU Alexa Fluor 555 Imaging Kit was
purchased from Thermo Fisher Scientific and manufacturer‟s protocol was followed with
minor modifications. Briefly, tissues were permeabilized in PBST for 20 minutes, post-
fixed in 4% PFA/PBST for ten minutes at room temperature, and then washed in PBST.
Tissues were incubated with EdU reagent for ten minutes, protected from light, at room
temperature. Thereafter, the tissues were washed thoroughly in PBST and prepared for
immunofluorescence.
97
β -Gal Antibody As Proxy For Robo Expression
After EdU labeling with AlexaFluor-555, slides were blocked with 3%BSA/PBST
for one hour at room temperature. Because Robo1 and Robo2 were replaced with LacZ
by knockin, lacZ transcript is transcribed from the Robo1 and Robo2 loci. The presence
of bGalactosidase protein, which is produced from the lacZ transcript, is accordingly an
appropriate proxy to show Robo expression domains. Thus, an antibody against β-
galactosidase was used as a proxy to mark Robo expression domains in the vertebral
column.
Primary antibody was diluted 1:200 in 3% BSA/PBST and tissues were incubated
overnight at 4° in a humidified slide box. A 488-conjugated secondary antibody was
selected for EdU quantification experiments. Secondary was incubated at one hour at
room temperature, protected from light. DAPI was used to counterstain nuclei and slides
were mounted using #1.5 glass coverslips and Dako fluorescent mounting medium.
Antibody information can be found in Table 1.
Quantification
Images were captured on a Leica confocal microscope. Effort was made to select
comparable regions from the trunk region containing Robo expressing tissues adjacent
to the notochord/disc. This was to ensure that quantification was based on proliferation
occurring in developmentally congruent regions since the vertebral column develops
rostro-caudally and rostral tissue is developmentally more advanced than caudal tissue.
Leica software was used to separate fluorescent images into separate channels
to quantify EdU-labeled and DAPI-labeled nuclei. Only nuclei within b-galactosidase-
488 labeled regions were counted. Quantification was done in a double-blind fashion. A
98
numerical code was assigned to all images to conceal genotype information and a
research volunteer in the lab was asked to manually count DAPI and EdU labeled
nuclei.
One midline sagittal image from each embryo was used for quantification. Each
section contained 2-5 well-defined Robo expressing domains (by bGalatosidase-488
proxy), and these were used to define regions for quantification. Total number of EdU-
555 positive nuclei within the 488-positive domains was compared to total number of
DAPI labeled nuclei within the same 488-positive domains to calculate % proliferation in
each embryo. Three Robo1+/-;Robo2+/- controls and three Robo1-/-;Robo2-/- embryos
were used.
Percent proliferation in the notochord/discs, which do not express Robo and are
thus negative for bGalatosidase-488, was also calculated as a control for any potential
changes in overall mitotic index.
Growth Plate Analysis
Markers for growth plate chondrocyte organization were evaluated using
antisense RNA probes to Col10a1 and Ihh, which are expressed by hypertrophic and
pre-hypertrophic chondrocytes, respectively13. In situ hybridizations were performed as
described above.
Characterization of Wnt/β-Catenin Signaling Activity
TCF/Lef:H2B-GFP reporter mice were used to evaluate Wnt/β-Catenin signaling
activity in the notochord, somites, and postnatal intervertebral discs and vertebrae.
Tissues were fixed overnight 4%PFA/PBS at 4°, cryopreserved in 30% sucrose
overnight, embedded in OCT embedding medium, and sectioned on a cryostat.
99
Postnatal tissues were additionally de-calcified overnight in 0.5M EDTA (pH 8) at 4°
before cryopreservation in 30% sucrose and embedding in OCT embedding medium. All
vertebral columns were sectioned through the sagittal plane at 20um or 40um. To
visualize GFP reporter activity, tissue sections were permeabilized in 0.1% PBST (0.1%
Tween-20 in PBS) and then counterstained with DAPI to show tissue architecture.
Tissues were mounted with DAKO fluorescent mounting media, and imaged on a Leica
confocal microscope.
Table 4-1. ECM Immunofluorscence And Western Blot Antibodies
Immunofluorescence Primary Antibodies
Aggrecan Millipore AB1031: polyclonal antibody against mouse aggrecan AA1177-1326
β-galactosidase Abcam: ab9361
Col1a1 Santa Cruz (D-13): sc-25974
Col2a1 Santa Cruz (C-19): sc-7763
Pax1 Abcam: ab95227
Slit2 GeneTex (FLJ14420): GTX118220
Tenascin C Santa Cruz (H-300): sc-20932
Tenascin C GeneTex (EPR4219): GTX62552
Tenascin C GeneTex (MTn-12): GTX26346)
T-brachyury Santa Cruz (N-19): sc-17743
Western blot primary antibodies
Tenascin C Santa Cruz (H-300): sc-20932
GAPDH Abcam (6C5): ab8245
Table 4-2. Genotyping Primers
Primers References
Robo1Null FwdCommon: TGGCACGAAGGTATATGTGC RevWT: GAAGGACTGGTGGTTTTGAG RevNull: CCTCCGCAAACTCCTATTTC
Long, et al.
Robo2Null FwdCommon: AAGTGCAACGTCTCTGAAGTCCC RevWT: GGCGGAATTCTTAATTAAGGCGCG RevNull: TTCTTTAGAAGGCACAACAATCTCAGAG
Grieshammer, et al.
100
Robo2FLOXED FwdFloxed: CCAATCATAGTCTCTCCACG RevFloxed: CCTCTGATTCAATGAGATGC
Lu, et al.
Rosa-mT/mG Fwd: CTCTGCTGCCTCCTGGCTTCT RevWT: CGAGGCGGATCACAAGCAATA RevMTMG: TCAATGGGCGGGGGTCGTT
Mouse generated by Muzumdar, et al.
Allele Primers References
TCF4-GFP FwdGFP: ACAACAAGCGCTCGACCATCAC RevGFP: AGTCGATGCCCTTCAGCTCGAT
Mouse generated by Ferrer-Vaquer, et al.
101
REFERENCES
Abbas J, Hamoud K, May H, Peled N, Sarig R, Stein D, Alperovitch-Najemson D, Hershkovitz I. Socioeconomic and physical characteristics of individuals with degenerative lumbar spinal stenosis. Spine (Phila Pa 1976) 2013, 38:E554-561.
Abdelkhalek HB, Beckers A, Schuster-Gossler K, Pavlova MN, Burkhardt H, Lickert H, Rossant J, Reinhardt R, Schalkwyk LC, Muller I, et al. The mouse homeobox gene Not is required for caudal notochord development and affected by the truncate mutation. Genes Dev 2004, 18:1725-1736.
Adham IM, Gille M, Gamel AJ, Reis A, Dressel R, Steding G, Brand-Saberi B, Engel W. The scoliosis (sco) mouse: a new allele of Pax1. Cytogenet Genome Res 2005, 111:16-26.
Akazawa H, Komuro I, Sugitani Y, Yazaki Y, Nagai R, Noda T. Targeted disruption of the homeobox transcription factor Bapx1 results in lethal skeletal dysplasia with asplenia and gastroduodenal malformation. Genes Cells 2000, 5:499-513.
Akiyama H, Chaboissier MC, Martin JF, Schedl A, de Crombrugghe B. 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 2002, 16:2813-2828.
Andersson GB. Epidemiological features of chronic low-back pain. Lancet 1999, 354:581-585.
Ang SL, Rossant J. HNF-3 beta is essential for node and notochord formation in mouse development. Cell 1994, 78:561-574.
Aoyama H, Asamoto K. The developmental fate of the rostral/caudal half of a somite for vertebra and rib formation: experimental confirmation of the resegmentation theory using chick-quail chimeras. Mech Dev 2000, 99:71-82.
Aoyama, H., Mizutani-koseki, S., Koseki, H. (2005) Three developmental compartments involved in rib formation. Int J Dev Biol. 49(2-3): 325-33.
Baffi MO, Moran MA, Serra R. Tgfbr2 regulates the maintenance of boundaries in the axial skeleton. Dev Biol 2006, 296:363-374.
Baffi MO, Slattery E, Sohn P, Moses HL, Chytil A, Serra R. Conditional deletion of the TGF-beta type II receptor in Col2a expressing cells results in defects in the axial skeleton without alterations in chondrocyte differentiation or embryonic development of long bones. Dev Biol 2004, 276:124-142.
102
Bagnall KM. The migration and distribution of somite cells after labelling with the carbocyanine dye, Dil: the relationship of this distribution to segmentation in the vertebrate body. Anat Embryol (Berl) 1992, 185:317-324.
Barrionuevo F, Taketo MM, Scherer G, Kispert A. Sox9 is required for notochord maintenance in mice. Dev Biol 2006, 295:128-140.
Beddington RS, Rashbass P, Wilson V. Brachyury--a gene affecting mouse gastrulation and early organogenesis. Dev Suppl 1992:157-165.
Bi W, Deng JM, Zhang Z, Behringer RR, de Crombrugghe B. Sox9 is required for cartilage formation. Nat Genet 1999, 22:85-89.
Blockus, H, Chedotal, A. Slit-Robo signaling. Development, 2016, 143(17):3037-44.
Borycki AG, Brunk B, Tajbakhsh S, Buckingham M, Chiang C, Emerson CP, Jr. Sonic hedgehog controls epaxial muscle determination through Myf5 activation. Development 1999, 126:4053-4063.
Brent AE, Braun T, Tabin CJ. Genetic analysis of interactions between the somitic muscle, cartilage and tendon cell lineages during mouse development. Development 2005, 132:515-528.
Brent AE, Schweitzer R, Tabin CJ. A somitic compartment of tendon progenitors. Cell 2003, 113:235-248.
Brent AE, Tabin CJ. FGF acts directly on the somitic tendon progenitors through the Ets transcription factors Pea3 and Erm to regulate scleraxis expression. Development 2004, 131:3885-3896.
Bruggeman BJ, Maier JA, Mohiuddin YS, Powers R, Lo Y, Guimaraes-Camboa N, Evans SM, Harfe BD. Avian intervertebral disc arises from rostral sclerotome and lacks a nucleus pulposus: implications for evolution of the vertebrate disc. Dev Dyn 2012, 241:675-683.
Buchbinder R, Blyth FM, March LM, Brooks P, Woolf AD, Hoy DG. Placing the global burden of low back pain in context. Best Pract Res Clin Rheumatol 2013, 27:575-589.
Bussen M, Petry M, Schuster-Gossler K, Leitges M, Gossler A, Kispert A. The T-box transcription factor Tbx18 maintains the separation of anterior and posterior somite compartments. Genes Dev 2004, 18:1209-1221.
Buttitta L, Mo R, Hui CC, Fan CM. Interplays of Gli2 and Gli3 and their requirement in mediating Shh-dependent sclerotome induction. Development 2003, 130:6233-6243.
103
Cadigan, K.M., Nusse, R. (1997) Wnt signaling: a common theme in animal development. Genes Dev. 11(24):3286-305.
Capellini TD, Zewdu R, Di Giacomo G, Asciutti S, Kugler JE, Di Gregorio A, Selleri L. Pbx1/Pbx2 govern axial skeletal development by controlling Polycomb and Hox in mesoderm and Pax1/Pax9 in sclerotome. Dev Biol 2008, 321:500-514.
Cerda J, Grund C, Franke WW, Brand M. Molecular characterization of Calymmin, a novel notochord sheath-associated extracellular matrix protein in the zebrafish embryo. Dev Dyn 2002, 224:200-209.
Chen, L., Liao, J., Klineberg, E., et al. Small leucine-rich proteoglycans (SLRPs): characteristics and function in the intervertebral disc. J Tissue Eng Regen Med, 2015.
Cheyette, B.N., Waxman, J.S., Miller, J.R., Takemaru, K., et al. (2002) Dapper, a Dishevelled-associated antagonist of beta-catenin and JNK signaling, is required for notochord formation. Dev Cell. 2(4): 449-61.
Chiang C, Litingtung Y, Lee E, Young KE, Corden JL, Westphal H, Beachy PA. Cyclopia and defective axial patterning in mice lacking Sonic hedgehog gene function. Nature 1996, 383:407-413.
Chiquet-Ehrismann, R., Kalla, P., Pearson, C., et al. (1988) Tenascin interferes with fibronectin action. Cell. 53(3): 383-390.
Choi KS, Harfe BD. Hedgehog signaling is required for formation of the notochord sheath and patterning of nuclei pulposi within the intervertebral discs. Proc Natl Acad Sci U S A 2011, 108:9484-9489.
Choi KS, Lee C, Harfe BD. Sonic hedgehog in the notochord is sufficient for patterning of the intervertebral discs. Mech Dev 2012, 129:255-262.
Christ B, Huang R, Scaal M. Formation and differentiation of the avian sclerotome. Anat Embryol (Berl) 2004, 208:333-350.
Christ B, Scaal M. Formation and differentiation of avian somite derivatives. Adv Exp Med Biol 2008, 638:1-41.
Cleaver O, Krieg PA. Notochord patterning of the endoderm. Dev Biol 2001, 234:1-12.
Colombier P, Clouet J, Hamel O, Lescaudron L, Guicheux J. The lumbar intervertebral disc: from embryonic development to degeneration. Joint Bone Spine 2014, 81:125-129.
104
Corallo D, Trapani V, Bonaldo P. The notochord: structure and functions. Cell Mol Life Sci 2015.
Cortes DH, Elliott DM. Extra-fibrillar matrix mechanics of annulus fibrosus in tension and compression. Biomech Model Mechanobiol 2012, 11:781-790.
Cserjesi P, Brown D, Ligon KL, Lyons GE, Copeland NG, Gilbert DJ, Jenkins NA, Olson EN. Scleraxis: a basic helix-loop-helix protein that prefigures skeletal formation during mouse embryogenesis. Development 1995, 121:1099-1110.
Dahia CL, Mahoney E, Wylie C. Shh signaling from the nucleus pulposus is required for the postnatal growth and differentiation of the mouse intervertebral disc. PLoS One 2012, 7:e35944.
Dahia CL, Mahoney EJ, Durrani AA, C. Intercellular signaling pathways active during intervertebral disc growth, differentiation, and aging. Spine (Phila Pa 1976) 2009, 34:456-462.
Dario AB, Ferreira ML, Refshauge KM, Lima TS, Ordonana JR, Ferreira PH. The relationship between obesity, low back pain, and lumbar disc degeneration when genetics and the environment are considered: a systematic review of twin studies. Spine J 2015, 15:1106-1117.
DasGupta, R., Fuchs, E. (1999) Multiple roles for activated LEF/TCF transcription complexes during hair follicle development and differentiation. Development. 126(20):4557-68.
Dawes B. The development of the vertebral column in mammals, as illustrated by its development in Mus musculus. Philosophical Transactions of the Royal Society of London Series B-Containing Papers of a Biological Character 1930, 218:115-U122.
de Crombrugghe B, Lefebvre V, Behringer RR, Bi W, Murakami S, Huang W. Transcriptional mechanisms of chondrocyte differentiation. Matrix Biol 2000, 19:389-394.
Deries M, Schweitzer R, Duxson MJ. Developmental fate of the mammalian myotome. Dev Dyn 2010, 239:2898-2910.
Deutsch U, Dressler GR, Gruss P. Pax 1, a member of a paired box homologous murine gene family, is expressed in segmented structures during development. Cell 1988, 53:617-625.
DiPaola CP, Farmer JC, Manova K, Niswander LA. Molecular signaling in intervertebral disk development. J Orthop Res 2005, 23:1112-1119.
105
Ewan KB, Everett AW. Evidence for resegmentation in the formation of the vertebral column using the novel approach of retroviral-mediated gene transfer. Exp Cell Res 1992, 198:315-320.
Fan CM, Lee CS, Tessier-Lavigne M. A role for WNT proteins in induction of dermomyotome. Dev Biol 1997, 191:160-165.
Fan CM, Tessier-Lavigne M. Patterning of mammalian somites by surface ectoderm and notochord: evidence for sclerotome induction by a hedgehog homolog. Cell 1994, 79:1175-1186.
Freemont AJ, Watkins A, Le Maitre C, Baird P, Jeziorska M, Knight MT, Ross ER, O'Brien JP, Hoyland JA. Nerve growth factor expression and innervation of the painful intervertebral disc. J Pathol 2002, 197:286-292.
Gotz W, Osmers R, Herken R. Localisation of extracellular matrix components in the embryonic human notochord and axial mesenchyme. J Anat 1995, 186 ( Pt 1):111-121.
Goulding MD, Chalepakis G, Deutsch U, Erselius JR, Gruss P. Pax-3, a novel murine DNA binding protein expressed during early neurogenesis. EMBO J 1991, 10:1135-1147.
Gruber HE, Ingram JA, Davis DE, Hanley EN, Jr. Increased cell senescence is associated with decreased cell proliferation in vivo in the degenerating human annulus. Spine J 2009, 9:210-215.
Gruber HE, Ingram JA, Norton HJ, Hanley EN, Jr. Senescence in cells of the aging and degenerating intervertebral disc: immunolocalization of senescence-associated beta-galactosidase in human and sand rat discs. Spine (Phila Pa 1976) 2007, 32:321-327.
Gruneberg H. Genetical studies on the skeleton of the mouse. XXIII. The development of brachyury and anury. J Embryol Exp Morphol 1958, 6:424-443.
Haque M, Ohata K, Takami T, Soares SB, Jr., Aree SN, Hakuba A, Hara M. Development of lumbosacral spina bifida: three-dimensional computer graphic study of human embryos at Carnegie stage twelve. Pediatr Neurosurg 2001, 35:247-252.
Harfe BD, Scherz PJ, Nissim S, Tian H, McMahon AP, Tabin CJ. Evidence for an expansion-based temporal Shh gradient in specifying vertebrate digit identities. Cell 2004, 118:517-528.
106
Hargus G, Kist R, Kramer J, Gerstel D, Neitz A, Scherer G, Rohwedel J. Loss of Sox9 function results in defective chondrocyte differentiation of mouse embryonic stem cells in vitro. Int J Dev Biol 2008, 52:323-332.
Hay ED. The mesenchymal cell, its role in the embryo, and the remarkable signaling mechanisms that create it. Dev Dyn 2005, 233:706-720.
Hayes AJ, Isaacs MD, Hughes C, Caterson B, Ralphs JR. Collagen fibrillogenesis in the development of the annulus fibrosus of the intervertebral disc. Eur Cell Mater 2011, 22:226-241.
Henry SP, Liang S, Akdemir KC, de Crombrugghe B. The postnatal role of Sox9 in cartilage. J Bone Miner Res 2012, 27:2511-2525.
Herbrand H, Pabst O, Hill R, Arnold HH. Transcription factors Nkx3.1 and Nkx3.2 (Bapx1) play an overlapping role in sclerotomal development of the mouse. Mech Dev 2002, 117:217-224.
Herrmann BG. Action of the Brachyury gene in mouse embryogenesis. Ciba Found Symp 1992, 165:78-86; discussion 86-91.
Hestbaek L, Iachine IA, Leboeuf-Yde C, Kyvik KO, Manniche C. Heredity of low back pain in a young population: a classical twin study. Twin Res 2004, 7:16-26.
Hiyama, A., Sakai, D., Risbud, M.V., et al. (2013) Enhancement of Intervertebral Disc Cell Senescence by Wnt/β-Catenin Signaling-Induced Matrix Metalloproteinase Expression. Arthritis Rheum. 62(10):3036-3047.
Hohenester, E. Structural insight into Slit-Robo signaling. Biochem Soc Trans, 2008,
(36):251-6. Holmes, G., Niswander, L. Expression of slit-2 and slit-3 during chick development. Dev
Dyn, 2001, 222(2):301-7. Howitt, J.A., Clout, N.J., Hohenester, E. Binding site for Robo receptors revealed by
dissection of the leucine-rich repeat region of Slit. EMBO J, 2004, (22):4406-12.
Hui CC, Joyner AL. A mouse model of greig cephalopolysyndactyly syndrome: the extra-toesJ mutation contains an intragenic deletion of the Gli3 gene. Nat Genet 1993, 3:241-246.
Huang, W., Chiquet-Ehrismann, R., Moyano, J.V., et al. (2001). Interference of Tenascin-C with Syndecan-4 Binding to Fibronectin Blocks Cell Adhesion and Stimulates Tumor Cell Proliferation. Cancer Res. 61(23): 8586-94.
107
Ikeda T, Kawaguchi H, Kamekura S, Ogata N, Mori Y, Nakamura K, Ikegawa S, Chung UI. Distinct roles of Sox5, Sox6, and Sox9 in different stages of chondrogenic differentiation. J Bone Miner Metab 2005, 23:337-340.
Jeong Y, Epstein DJ. Distinct regulators of Shh transcription in the floor plate and notochord indicate separate origins for these tissues in the mouse node. Development 2003, 130:3891-3902.
Johnson ZI, Shapiro IM, Risbud MV. Extracellular osmolarity regulates matrix homeostasis in the intervertebral disc and articular cartilage: evolving role of TonEBP. Matrix Biol 2014, 40:10-16.
Jostes B, Walther C, Gruss P. The murine paired box gene, Pax7, is expressed specifically during the development of the nervous and muscular system. Mech Dev 1990, 33:27-37.
Kaestner KH, Hiemisch H, Luckow B, Schutz G. The HNF-3 gene family of transcription factors in mice: gene structure, cDNA sequence, and mRNA distribution. Genomics 1994, 20:377-385.
Katz JN. Lumbar disc disorders and low-back pain: socioeconomic factors and consequences. J Bone Joint Surg Am 2006, 88 Suppl 2:21-24.
Kiani C, Chen L, Wu YJ, Yee AJ, Yang BB. Structure and function of aggrecan. Cell Res 2002, 12:19-32.
Kim, M., Fontelonga, T., Roesener, A.P., Lee, H., Gurung, S., et al. Motor neuron cell bodies are actively positioned by Slit/Robo repulsion and Netrin/DCC attraction. Dev Biol, 2015. 399(1):68-79.
Kimmel CB, Ballard WW, Kimmel SR, Ullmann B, Schilling TF. Stages of embryonic development of the zebrafish. Dev Dyn 1995, 203:253-310.
Koseki H, Wallin J, Wilting J, Mizutani Y, Kispert A, Ebensperger C, Herrmann BG, Christ B, Balling R. A role for Pax-1 as a mediator of notochordal signals during the dorsoventral specification of vertebrae. Development 1993, 119:649-660.
Kraus F, Haenig B, Kispert A. Cloning and expression analysis of the mouse T-box gene Tbx18. Mech Dev 2001, 100:83-86.
Lefebvre, V., Li, P., de Crombrugghe, B. (1998) A new long form of Sox5 (L-Sox5), Sox6 and Sox9 are coexpressed in chondrogenesis and cooperatively activate the type II collagen gene. EMBO J. 17(19):5718-5733.
108
Le Maitre CL, Freemont AJ, Hoyland JA. Accelerated cellular senescence in degenerate intervertebral discs: a possible role in the pathogenesis of intervertebral disc degeneration. Arthritis Res Ther 2007, 9:R45.
Lehtonen E, Stefanovic V, Saraga-Babic M. Changes in the expression of intermediate filaments and desmoplakins during development of human notochord. Differentiation 1995, 59:43-49.
Leitges M, Neidhardt L, Haenig B, Herrmann BG, Kispert A. The paired homeobox gene Uncx4.1 specifies pedicles, transverse processes and proximal ribs of the vertebral column. Development 2000, 127:2259-2267.
Lettice L, Hecksher-Sorensen J, Hill R. The role of Bapx1 (Nkx3.2) in the development and evolution of the axial skeleton. J Anat 2001, 199:181-187.
Li Y, Litingtung Y, Ten Dijke P, Chiang C. Aberrant Bmp signaling and notochord delamination in the pathogenesis of esophageal atresia. Dev Dyn 2007, 236:746-754.
Liu H, Zhang C, Zhu S, Lu P, Zhu T, Gong X, Zhang Z, Hu J, Yin Z, Heng BC, et al. Mohawk promotes the tenogenesis of mesenchymal stem cells through activation of the TGFbeta signaling pathway. Stem Cells 2015, 33:443-455.
Lv, F.J., Peng, Y., Sun, Y., et al. (2016) Matrix metalloproteinase 12 is an indicator of intervertebral disc degeneration co-expressed with fibrotic markers. Osteoarthritis Cartilage. 24(10):1826-36.
Maier JA, Harfe BD. Nuclei pulposi formation from the embryonic notochord occurs normally in GDF-5-deficient mice. Spine (Phila Pa 1976) 2011, 36:E1555-1561.
Maier JA, Lo Y, Harfe BD. Foxa1 and Foxa2 are required for formation of the intervertebral discs. PLoS One 2013, 8:e55528.
Malandrino A, Lacroix D, Hellmich C, Ito K, Ferguson SJ, Noailly J. The role of endplate poromechanical properties on the nutrient availability in the intervertebral disc. Osteoarthritis Cartilage 2014, 22:1053-1060.
Mansouri A, Yokota Y, Wehr R, Copeland NG, Jenkins NA, Gruss P. Paired-related murine homeobox gene expressed in the developing sclerotome, kidney, and nervous system. Dev Dyn 1997, 210:53-65.
MacDonald, B.T., Tamai, K., He, X. (2009) Wnt/beta-catenin signaling: components, mechanisms, and diseases. Dev Cell. 17(1): 9-26.
109
McCann MR, Tamplin OJ, Rossant J, Seguin CA. Tracing notochord-derived cells using a Noto-cre mouse: implications for intervertebral disc development. Dis Model Mech 2012, 5:73-82.
McMahon JA, Takada S, Zimmerman LB, Fan CM, Harland RM, McMahon AP. Noggin-mediated antagonism of BMP signaling is required for growth and patterning of the neural tube and somite. Genes Dev 1998, 12:1438-1452.
Merceron C, Mangiavini L, Robling A, Wilson TL, Giaccia AJ, Shapiro IM, Schipani E, Risbud MV. Loss of HIF-1alpha in the notochord results in cell death and complete disappearance of the nucleus pulposus. PLoS One 2014, 9:e110768.
Millennium WHOSGotBoMCatSotN. The burden of musculoskeletal conditions at the start of the new millennium. World Health Organ Tech Rep Ser 2003, 919:i-x, 1-218, back cover.
Mo R, Freer AM, Zinyk DL, Crackower MA, Michaud J, Heng HH, Chik KW, Shi XM, Tsui LC, Cheng SH, et al. Specific and redundant functions of Gli2 and Gli3 zinc finger genes in skeletal patterning and development. Development 1997, 124:113-123.
Murchison ND, Price BA, Conner DA, Keene DR, Olson EN, Tabin CJ, Schweitzer R. Regulation of tendon differentiation by scleraxis distinguishes force-transmitting tendons from muscle-anchoring tendons. Development 2007, 134:2697-2708.
Nasto LA, Ngo K, Leme AS, Robinson AR, Dong Q, Roughley P, Usas A, Sowa GA, Pola E, Kang J, et al. Investigating the role of DNA damage in tobacco smoking-induced spine degeneration. Spine J 2014, 14:416-423.
Neubuser A, Koseki H, Balling R. Characterization and developmental expression of Pax9, a paired-box-containing gene related to Pax1. Dev Biol 1995, 170:701-716.
Noel, L.S., Champion, B.R., Holley, C.L., et al. (1998) RoBo-1, a novel member of the urokinase plasminogen activator receptor/CD59/Ly-6/snake toxin family selectively expressed in rat bone and growth plate cartilage. J Biol Chem. 273(7):3878-83.
Olivera-Martinez I, Thelu J, Teillet MA, Dhouailly D. Dorsal dermis development depends on a signal from the dorsal neural tube, which can be substituted by Wnt-1. Mech Dev 2001, 100:233-244.
Ordan, E., Brankatschk, M., Dickson, B., et al. Slit cleavage is essential for producing an active, stable, non-diffusible short-range signal that guides muscle migration. Development, 2015. 142(8):1431-6.
110
Paavola LG, Wilson DB, Center EM. Histochemistry of the developing notochord, perichordal sheath and vertebrae in Danforth's short-tail (sd) and normal C57BL/6 mice. J Embryol Exp Morphol 1980, 55:227-245.
Peters H, Neubuser A, Kratochwil K, Balling R. Pax9-deficient mice lack pharyngeal pouch derivatives and teeth and exhibit craniofacial and limb abnormalities. Genes Dev 1998, 12:2735-2747.
Peters H, Wilm B, Sakai N, Imai K, Maas R, Balling R. Pax1 and Pax9 synergistically regulate vertebral column development. Development 1999, 126:5399-5408.
Pryce BA, Watson SS, Murchison ND, Staverosky JA, Dunker N, Schweitzer R. Recruitment and maintenance of tendon progenitors by TGFbeta signaling are essential for tendon formation. Development 2009, 136:1351-1361.
Rashbass P, Wilson V, Rosen B, Beddington RS. Alterations in gene expression during mesoderm formation and axial patterning in Brachyury (T) embryos. Int J Dev Biol 1994, 38:35-44.
Richardson SM, Purmessur D, Baird P, Probyn B, Freemont AJ, Hoyland JA. Degenerate human nucleus pulposus cells promote neurite outgrowth in neural cells. PLoS One 2012, 7:e47735.
Riddle RD, Johnson RL, Laufer E, Tabin C. Sonic hedgehog mediates the polarizing activity of the ZPA. Cell 1993, 75:1401-1416.
Rodrigo I, Hill RE, Balling R, Munsterberg A, Imai K. Pax1 and Pax9 activate Bapx1 to induce chondrogenic differentiation in the sclerotome. Development 2003, 130:473-482.
Romgens AM, van Donkelaar CC, Ito K. Contribution of collagen fibers to the compressive stiffness of cartilaginous tissues. Biomech Model Mechanobiol 2013, 12:1221-1231.
Roussa E, Krieglstein K. Induction and specification of midbrain dopaminergic cells: focus on SHH, FGF8, and TGF-beta. Cell Tissue Res 2004, 318:23-33.
Saga, Y. The mechanism of somite formation in mice. Curr Opin Genet Dev 2012, 22(4):331-8.
Saraga-Babic M, Lehtonen E, Svajger A, Wartiovaara J. Morphological and immunohistochemical characteristics of axial structures in the transitory human tail. Ann Anat 1994, 176:277-286.
111
Schneider PR, Buhrmann C, Mobasheri A, Matis U, Shakibaei M. Three-dimensional high-density co-culture with primary tenocytes induces tenogenic differentiation in mesenchymal stem cells. J Orthop Res 2011, 29:1351-1360.
Selleri L, Depew MJ, Jacobs Y, Chanda SK, Tsang KY, Cheah KS, Rubenstein JL, O'Gorman S, Cleary ML. Requirement for Pbx1 in skeletal patterning and programming chondrocyte proliferation and differentiation. Development 2001, 128:3543-3557.
Senthinathan B, Sousa C, Tannahill D, Keynes R. The generation of vertebral segmental patterning in the chick embryo. J Anat 2012, 220:591-602.
Shukunami C, Takimoto A, Oro M, Hiraki Y. Scleraxis positively regulates the expression of tenomodulin, a differentiation marker of tenocytes. Dev Biol 2006, 298:234-247.
Sivan SS, Hayes AJ, Wachtel E, Caterson B, Merkher Y, Maroudas A, Brown S, Roberts S. Biochemical composition and turnover of the extracellular matrix of the normal and degenerate intervertebral disc. Eur Spine J 2014, 23 Suppl 3:S344-353.
Sivan SS, Wachtel E, Roughley P. Structure, function, aging and turnover of aggrecan in the intervertebral disc. Biochim Biophys Acta 2014, 1840:3181-3189.
Smith LJ, Elliott DM. Formation of lamellar cross bridges in the annulus fibrosus of the intervertebral disc is a consequence of vascular regression. Matrix Biol 2011, 30:267-274.
Smith LJ, Nerurkar NL, Choi KS, Harfe BD, Elliott DM. Degeneration and regeneration of the intervertebral disc: lessons from development. Dis Model Mech 2011, 4:31-41.
Smits P, Lefebvre V. Sox5 and Sox6 are required for notochord extracellular matrix sheath formation, notochord cell survival and development of the nucleus pulposus of intervertebral discs. Development 2003, 130:1135-1148.
Smits P, Li P, Mandel J, Zhang Z, Deng JM, Behringer RR, de Crombrugghe B, Lefebvre V. The transcription factors L-Sox5 and Sox6 are essential for cartilage formation. Dev Cell 2001, 1:277-290.
Smolders, L. A., Meij, B. P., Onis, D., Riemers, F. M., et al. (2013) Gene expression profiling of early intervertebral disc degeneration reveals a down-regulation of canonical Wnt signaling and caveolin-1 expression: implications for development of regenerative strategies. Arthritis Res Ther. 15(1):R23.
112
Sofat, N., Robertson, S.D., Hermansson, M., et al. (2012) Tenascin-C fragments are endogenous inducers of cartilage matrix degradation. Rheumatol Int. 32(9):2809-2817.
Stemple DL. Structure and function of the notochord: an essential organ for chordate development. Development 2005, 132:2503-2512.
Sugimoto Y, Takimoto A, Akiyama H, Kist R, Scherer G, Nakamura T, Hiraki Y, Shukunami C. Scx+/Sox9+ progenitors contribute to the establishment of the junction between cartilage and tendon/ligament. Development 2013, 140:2280-2288.
Sulik K, Dehart DB, Iangaki T, Carson JL, Vrablic T, Gesteland K, Schoenwolf GC. Morphogenesis of the murine node and notochordal plate. Dev Dyn 1994, 201:260-278.
Takahashi Y, Yasuhiko Y, Takahashi J, Takada S, Johnson RL, Saga Y, Kanno J. Metameric pattern of intervertebral disc/vertebral body is generated independently of Mesp2/Ripply-mediated rostro-caudal patterning of somites in the mouse embryo. Dev Biol 2013, 380:172-184.
Takimoto A, Mohri H, Kokubu C, Hiraki Y, Shukunami C. Pax1 acts as a negative regulator of chondrocyte maturation. Exp Cell Res 2013, 319:3128-3139.
Takimoto A, Oro M, Hiraki Y, Shukunami C. Direct conversion of tenocytes into chondrocytes by Sox9. Exp Cell Res 2012, 318:1492-1507.
Talbot, W.S., Trevarrow, B., Halpern, M.E., et al. (1995) A homeobox gene essential for zebrafish notochord development. Nature. 378(6553):150-7.
Tam PP, Trainor PA. Specification and segmentation of the paraxial mesoderm. Anat Embryol (Berl) 1994, 189:275-305.
Tamplin OJ, Cox BJ, Rossant J. Integrated microarray and ChIP analysis identifies multiple Foxa2 dependent target genes in the notochord. Dev Biol 2011, 360:415-425.
Tanaka M, Komuro I, Inagaki H, Jenkins NA, Copeland NG, Izumo S. Nkx3.1, a murine homolog of Ddrosophila bagpipe, regulates epithelial ductal branching and proliferation of the prostate and palatine glands. Dev Dyn 2000, 219:248-260.
Theiler K. Anatomy and development of the "truncate" (boneless) mutation in the mouse. Am J Anat 1959, 104:319-343.
Theiler K. The house mouse; development and normal stages from fertilization to 4 weeks of age. Berlin, New York,: Springer-Verlag; 1972.
113
Tolofari SK, Richardson SM, Freemont AJ, Hoyland JA. Expression of semaphorin 3A and its receptors in the human intervertebral disc: potential role in regulating neural ingrowth in the degenerate intervertebral disc. Arthritis Res Ther 2010, 12:R1.
Tribioli C, Lufkin T. The murine Bapx1 homeobox gene plays a critical role in embryonic development of the axial skeleton and spleen. Development 1999, 126:5699-5711.
Tylzanowski P, Mebis L, Luyten FP. The Noggin null mouse phenotype is strain dependent and haploinsufficiency leads to skeletal defects. Dev Dyn 2006, 235:1599-1607.
Ukita, K., Hirahara, S., Oshima, N., Imuta, Y., et al. (2009) Wnt signaling maintains the notochord fate for progenitor cells and supports the posterior extension of the notochord. Mech Dev. 126(10):791-803.
Urban, J., Roberts, S. (2003) Degeneration of the intervertebral disc. Arthritis Res Ther.
5(3):120-130. Usami, Y., Gunawardena, A.T., Iwamoto, M., Enomoto-Iwamoto, M. (2016). Wnt
signaling in cartilage development and diseases: lessons from animal studies. Lab Invest. 96(2): 186-96.
Wallin J, Wilting J, Koseki H, Fritsch R, Christ B, Balling R. The role of Pax-1 in axial skeleton development. Development 1994, 120:1109-1121.
Weinstein DC, Ruiz i Altaba A, Chen WS, Hoodless P, Prezioso VR, Jessell TM, Darnell JE, Jr. The winged-helix transcription factor HNF-3 beta is required for notochord development in the mouse embryo. Cell 1994, 78:575-588.
White TD, Black MT, Folkens PA. Human osteology. 3rd ed. San Diego, Calif.: Academic Press; 2012.
Wilkinson DG, Bhatt S, Herrmann BG. Expression pattern of the mouse T gene and its role in mesoderm formation. Nature 1990, 343:657-659.
Wilm B, Dahl E, Peters H, Balling R, Imai K. Targeted disruption of Pax1 defines its null phenotype and proves haploinsufficiency. Proc Natl Acad Sci U S A 1998, 95:8692-8697.
Winkler, T., Mahoney, E.J., Sinner, D., Wylie, C.C., Dahia, C.L. (2014) Wnt Signaling Activates Shh Signaling in Early Postnatal Intervertebral Discs, and Re-Activates Shh Signaling in Old Discs in the Mouse. PLoS One. 9(6):e98444.
114
Yamanaka Y, Tamplin OJ, Beckers A, Gossler A, Rossant J. Live imaging and genetic analysis of mouse notochord formation reveals regional morphogenetic mechanisms. Dev Cell 2007, 13:884-896.
Yamashita S, Andoh M, Ueno-Kudoh H, Sato T, Miyaki S, Asahara H. Sox9 directly promotes Bapx1 gene expression to repress Runx2 in chondrocytes. Exp Cell Res 2009, 315:2231-2240.
Yang, X.R., Ng, D., Alcorta, D.A., et al. (2009) T (Brachyury) gene duplication confers major susceptibility to familial chordoma. Nat Genet. 41(11):1176-8.
Ypsilanti, A.R., Zagar, Y., Chedotal, A. Moving away from the midline: new developments for Slit and Robo. Development, 2010. 137(12):1939-52.
Yusuf F, Brand-Saberi B. The eventful somite: patterning, fate determination and cell division in the somite. Anat Embryol (Berl) 2006, 211 Suppl 1:21-30.
Zhang XM, Ramalho-Santos M, McMahon AP. Smoothened mutants reveal redundant roles for Shh and Ihh signaling including regulation of L/R symmetry by the mouse node. Cell 2001, 106:781-792.
Zhao, J., Li, S., Trilok, S., et al. (2014) Small molecule-directed specification of sclerotome-like chondroprogenitors and induction of a somitic chondrogenesis program from embryonic stem cells. Development. 141(20):3848-58.
Zizic Mitrecic M, Mitrecic D, Pochet R, Kostovic-Knezevic L, Gajovic S. The mouse gene Noto is expressed in the tail bud and essential for its morphogenesis. Cells Tissues Organs 2010, 192:85-92.
115
BIOGRAPHICAL SKETCH
Lisa Lawson moved to the United States in 1990 from South Korea. She grew up
in Central Florida and graduated from Winter Park High School in 2003. She completed
her undergraduate studies at the University of Notre Dame from 2003-2007. After
completing her Bachelor of Science, she received a Master of Medical Sciences from
the University of South Florida. In 2009 she joined the Dickey lab at the University of
South Florida to work on Alzheimer‟s research, assisting in research on how autophagy
pathways can be manipulated to ameliorate tauopathies.In 2011, Lisa enrolled at the
University of Florida as a doctoral student to study vertebrate embryogenesis under the
tutelage of Brian Harfe.
