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Skeletal cell fate plasticity in zebrafish bone development and regeneration
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Skeletal cell fate plasticity in zebrafish bone development and regeneration
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Content
Skeletal Cell Fate Plasticity in Zebrafish Bone
Development and Regeneration
by
Dion F. Giovannone
A dissertation submitted in conformity with the requirements
for the degree of Doctor of Philosophy
in Development, Stem Cells, and Regenerative Medicine
FACULTY OF THE USC GRADUATE SCHOOL
University of Southern California
August 2019
© Copyright by Dion F. Giovannone 2019
ii
Acknowledgments
I would like to express my gratitude to my advisor, Dr. Gage Crump, for his counsel, advocacy,
and patience. His excellence in discerning thought, experimental design, and scientific
composition are ideals I will continue to strive towards as a scientific researcher. I consider
myself fortunate to have had a dedicated and exceptional mentor.
I am grateful for my committee members, Drs. Francesca Mariani and Amy Merrill, for their
constructive feedback, encouragement, and compassion.
I would like to offer special thanks to Drs. Lindsey Barske, Sandeep Paul, Joanna Smeeton, and
Camilla Teng for their generous instruction, experimental contributions, and enlightening
conversations concerning research objectives.
I would also like to acknowledge Simone Schindler and Punam Patel for superb technical
support and experimental troubleshooting; Megan Matsutani and Jennifer DeKoeyer Crump for
exceptional fish care; and members of the Crump lab for collaborative problem solving,
motivation, and optimism.
Finally, I dedicate my work to the memory of my parents for their immeasurable love and
inspiration. I am also deeply appreciative of my extended family, girlfriend, friends, and
classmates for their empathy, grace, and genuine care.
iii
Table of Contents
ACKNOWLEDGMENTS II
TABLE OF CONTENTS III
LIST OF FIGURES VI
CHAPTER 1 PROPERTIES OF ZEBRAFISH SKELETAL FORMATION AND REGENERATION 9
1 ABSTRACT 9
1.1 SKELETAL BONE DEVELOPMENT 9
1.1.1 ESTABLISHED SKELETAL CELL TYPES IN VERTEBRATES 10
1.1.2 CRANIOFACIAL SKELETAL BONE DIVERSITY IN ZEBRAFISH 12
1.1.3 PERIOSTEUM AND PERICHONDRIUM ROLES DURING BONE DEVELOPMENT 12
1.1.4 BONE REMODELING DURING GROWTH 13
1.2 CELL FATE PLASTICITY 14
1.2.1 CELL FATE PLASTICITY IN DEVELOPMENT AND INJURY 15
1.3 SKELETAL STEM AND PROGENITOR CELLS IN ADULTS 16
1.3.1 PERIOSTEUM AND PERICHONDRIUM ROLES DURING FRACTURE REPAIR 17
1.3.2 ADULT SKELETAL REGENERATION AND REPAIR IN ZEBRAFISH 17
1.4 REFERENCES 18
CHAPTER 2 PROGRAMMED CONVERSION OF HYPERTROPHIC CHONDROCYTES INTO OSTEOBLASTS
AND MARROW ADIPOCYTES WITHIN ZEBRAFISH BONES 26
2 ABSTRACT 26
2.1 INTRODUCTION 26
2.2 RESULTS 28
2.2.1 REMODELING OF THE CH BONE IN JUVENILE ZEBRAFISH 28
2.2.2 VASCULARIZATION OF THE CH BONE IN JUVENILE ZEBRAFISH 32
2.2.3 CONTRIBUTION OF SOX10-LINEAGE CELLS TO OSTEOBLASTS, ADIPOCYTES, AND MESENCHYMAL CELLS 34
iv
2.2.4 CONTRIBUTION OF COL2A1A+ CHONDROCYTES TO OSTEOBLASTS, ADIPOCYTES, AND
MESENCHYMAL CELLS 40
2.2.5 LONG-LIVED HISTONE2A-MCHERRY PROTEIN REVEALS CONTRIBUTION OF COL2A1A+
CHONDROCYTES TO OSTEOBLASTS 42
2.2.6 HYPERTROPHIC CHONDROCYTES RE-ENTER THE CELL CYCLE AND EXPRESS LEPR 47
2.2.7 REQUIREMENT OF MMP9 FOR GROWTH PLATE REMODELING AND MARROW ADIPOCYTE FORMATION 51
2.3 DISCUSSION 54
2.4 MATERIALS AND METHODS 57
2.4.1 ZEBRAfiSH TRANSGENIC LINES AND MMP9 MUTANTS 57
2.4.2 HISTOLOGY AND LIPIDTOX STAINING 58
2.4.3 IMMUNOHISTOCHEMISTRY AND IN SITU HYBRIDIZATION 59
2.4.4 NEURAL CREST TRANSPLANTATIONS 60
2.4.5 IMAGING 60
2.4.6 QUANTIFICATION AND STATISTICAL ANALYSES 60
2.5REFERENCES 61
CHAPTER 3 DEFINING THE PROGENITOR POPULATION IN ADULT ZEBRAFISH JAW BONE
REGENERATION 66
3 ABSTRACT 66
3.1 INTRODUCTION 66
3.2 RESULTS 70
3.2.1 LOWER JAWBONE REGENERATION PRODUCES A HYBRID CARTILAGE-BONE CALLUS 70
3.2.2 RUNX2+/SP7- PERIOSTEAL SUBPOPULATION GIVES RISE TO THE CARTILAGE CALLUS DURING JAWBONE
REGENERATION 73
3.2.3 JUVENILE LOWER JAW RESECTION 75
3.2.4 LINEAGE CONTRIBUTION OF OSTEOBLASTS AND CHONDROCYTES TO CARTILAGE CALLUS REGENERATION 77
3.2.5 UPPER JAWBONE REGENERATION 81
3.3 DISCUSSION 84
3.4 MATERIALS AND METHODS 85
3.4.1 HISTOLOGY AND PARAFFIN EMBEDDING 85
3.4.2 IMMUNOHISTOCHEMISTRY AND IN SITU HYBRIDIZATION 86
3.4.3 IMAGING 86
v
3.5 REFERENCES 87
CHAPTER 4 FUTURE PERSPECTIVES ABOUT SKELETAL CELL FATE PLASTICITY 91
4 ABSTRACT 91
4.1 ADIPOGENIC AND OSTEOGENIC CELL FATE DYNAMICS 91
4.2 INVADING IMMUNE AND VASCULATURE CELLS DYNAMICS WITH SKELETAL CELLS 93
4.3 OPPORTUNITIES TO DEFINE PERICHONDRAL AND PERIOSTEAL CELL POPULATIONS 94
4.4 REFERENCES 95
vi
List of Figures
Figure 2-1. Time-course of Ch remodeling in juvenile zebrafish 30
Figure supplement 2-1. Ch bone and marrow fat structure 31
Figure 2-2. Vascularization of the Ch 33
Figure 2-3. Contribution of sox10+ chondrocytes to osteoblasts and marrow
adipocytes 36
Figure supplement 2-2. Characterization of the sox10:CreERT2 and
col2a1a:CreERT2 transgenic lines 38
Figure supplement 2-3. Neural crest contributions to the Ch bone and marrow
adipocyte 39
Figure 2-4. Contribution of col2a1a+ chondrocytes to osteoblasts and marrow
adipocytes 41
Figure 2-5. Tracing of col2a1a-lineage cells by a long-lived Histone2A-mCherry
fusion protein 44
Figure supplement 2-4. Characterization of the col2a1a:Histone2A-mCherry-2a-
GFPCAAX line 46
Figure 2-6. Late-stage hypertrophic chondrocytes re-enter the cell cycle and
express lepr 49
Figure supplement 2-5. Expression of Lepr/lepr mRNA in zebrafish and mouse
endochondral bone 50
Figure 2-7. Tissue-autonomous requirement for mmp9 in cartilage remodeling 53
Figure 3-1. Working model of lower jawbone regeneration 70
Figure 3-2. Lower jawbone regeneration produces a hybrid cartilage-bone callus 72
Figure 3-3. Activation of the periosteum during jawbone regeneration 74
vii
Figure 3-4. Defining the quality of lower jawbone regeneration in juvenile fish 76
Figure supplement 3-1. Juvenile lower jawbone regeneration activates RUNX2+ cells
for cartilage callus formation 77
Figure 3-5. Contribution of derived osteoblasts and chondrocytes to cartilage callus
regeneration 80
Figure 3-6. Pre-maxilla resection regenerates in adult zebrafish 82
Figure supplement 3-2. Pre-maxilla regeneration generates cartilage while periosteal
cells co-express col2a1a and col1a1a 83
9
Chapter 1
Properties of Zebrafish skeletal formation and regeneration
1 Abstract
The skeletal system as an organ develops and regenerates through complex interactions with
surrounding tissues and vasculature. Cellular components of bone, cartilage, and fat all
demonstrate cell fate plasticity to contribute to the other skeletal cell lineages. Importantly, the
physical surroundings especially the building, breakdown, and remodeling of the matrix
components of bone, cartilage and fat are involved in the differentiation and dedifferentiation of
skeletal cells. While the periosteum is vital to fracture repair healing, the origins of and unique
molecular markers of the skeletal progenitor cell population are not yet understood. Using
zebrafish as a model for both skeletal development and regeneration can unravel the complex
interactions of many skeletal cell types while defining the induced or inherent skeletal cell fate
plasticity seen across vertebrates.
1.1 Skeletal Bone Development
The vertebrate skeleton develops through two main types of bone ossification, intramembranous
and endochondral ossification. Dermal bones are generated through intramembranous
ossification when mesenchymal cells condense and differentiate directly into osteoblasts
(Cunningham et al., 2016). In contrast, endochondral ossification initially utilizes a
mesenchymal condensation to differentiate into chondrocytes and produce a cartilage (Wagner
and Karsenty, 2001). However, a more complex third type of ossification, perichondral is
discussed during the mandibular bone formation around the Meckel’s cartilage (Weigele et al.
2016). There is a debate about if it should be categorized as either endochondral or
intramembranous ossification as it has characteristics of both types (Parada and Chai, 2016). In
both endochondral and perichondral, a cartilage template is used for further bone growth and
development. Since in mammals the Meckel’s cartilage is replaced by bone some classify it as
endochondral ossification. Others debate that since the perichondrium produce the osteoblasts to
10
generate the initial formation of a bone collar it is intramembranous ossification (Hall, 2016). In
zebrafish, the Meckel’s cartilage is maintained into adulthood and not replaced by bone at any
point during bone growth.
In endochondral bones, the simultaneous and coordinated growth in multiple axis and from
multiple centers including the radial growth from the periosteum is critical for not developing
Achondroplasias (Erlebacher et al., 1995). Notably the short wide bones of Achondroplasia are
also coupled with overgrowth of the periosteum. In fact, the osteogenic potential of the
periosteum reaches back to the 17
th
and 18
th
centuries (Bilkay et al., 2008; Ito et al., 2001). A
connection between the periosteum and cell signaling pathways was observed when Joyce et al.
(1990) demonstrated that TGFb influences differentiation of committed skeletal precursor cells
to generate cartilage and bone formation. Indications that the periosteum in mouse and newt
were involved in skeletal bone injury and regeneration reach back over 30 years (Neufeld et al.,
1985).
1.1.1 Established skeletal cell types in vertebrates
Chondrogenesis in early development is focused on the dynamic interactions of mesenchymal
stem cells undergoing condensations to establish the membranous skeleton. There are many open
questions about how the condensations are generated, especially on topics of spatiotemporal
skeletal cell migrations and recruitment of cells into the condensation (Widelitz et al., 1993;
Fang and Hall, 1999). The differentiation of the mesenchymal stem cells in the condensation into
chondroblasts and chondrocytes into the cartilage anlagen is the end result. Chondroblasts and
chondrocytes generate and deposit the cartilage ECM surrounding the cells. Chondroblasts
primarily proliferate to give rise to chondrocytes. Epithelial – mesenchymal interactions
involving cell adhesion molecules like NCAM and tenascin aid in the condensation transition
into differentiation (Hall, 2015).
Chondroblasts in mouse long bones are a highly proliferative cells that require the expression of
Sox5 and Sox6 to delay chondrocyte prehypertrophy (Smits et al., 2004). Further the induction
and formation of the prehypertrophic and hypertrophic zones involve Sox5/6 by delaying
terminal chondrocyte differentiation via affecting Ihh signaling, and Runx2 (Smits et al., 2004).
Additionally, Wnt7a can prevent chondrogenesis in chondrocytes and chondroblast
differentiation not only during condensation while working antagonistically against Bmp2 as
11
well as cartilage differentiation (Rudnicki and Brown, 1997; Staines et al., 2012; Ma et al., 2013;
Stott et al., 1999).
Osteoblasts and osteocytes are similar in cell type relation as chondroblasts are to chondrocytes.
Both osteoblasts and osteocytes deposit bone in the surrounding ECM. However, osteoblasts are
commonly observed on the surface of bone while osteocytes are located within mineralized bone
matrix. Osteoblasts are thought to produce a majority of the osteogenic bone matrix. Osteocytes
are osteoblasts that have ceased to divide and serve within the mineralized bone matrix as
mechano-sensory skeletal cells utilizing a system of Haversian canals and cellular processes
called canaliculi to form cellular junctions. There is debate about the origin of osteoclasts and
chondroclasts specifically are they monocyte derived, or macrophage derived (Hall, 2015).
Regardless they serve the primary function of reabsorption of bone or cartilage, usually but not
exclusively mineralized matrix. Both cell types can be mono- or multinucleated and osteoclasts
are functionally important for bone modeling and remodeling.
Adipose (fat) tissue is largely comprised of adipocytes, fat cells, and share a common
mesenchymal/mesodermal stem cell with osteogenic and chondrogenic lineages. Mouse Cre-
lineage tracing studies with conditional Sox9, Col2a1, and Aggrecan as transgenic labels have
demonstrated contribution to chondrocytes, osteoblasts, bone marrow, and adipocytes (Ono et
al., 2014). In fact, osteogenic signaling pathways like BMP-2, Wnt10b, and Hedgehog, work in
opposition to adipogenesis. The molecular mechanisms regulating adipogenesis is not fully
understood, mesenchymal stem cells that are adipogenic utilize Schnurri-2 to modify BMP-2
working in cooperation with Smad1, 4 and C/EBPa (Jin, 2006). Adipogenesis appears to focus
around three major families of transcription factors, C/EBP’s, PPAR-g, and KLF’s (Rosen and
MacDougald, 2006).
Mammals have brown and white adipose tissue while bony fishes have only been observed to
have white adipose tissue. White adipose tissue can be deposited in variety of locations in the
body and is thought to mainly serve as metabolic energy stores, while brown adipose tissue may
also serve in thermoregulation (Gesta et al., 2007). More recently adipose tissue has been shown
to be involved in endocrine function (Cristancho and Lazar, 2011). The deposition of adipocytes
within the bone marrow as a skeletal function is less understood. Perhaps the softness of marrow
adipocytes can absorb physical force stresses on bones that are constantly moving.
12
Adipocytes are identified primarily by a single lipid droplet that within the bone marrow of
zebrafish are uniquely larger than visceral adipocytes found outside bone marrow. Additionally,
Perilipin, C/EBP, and PPAR-g are established molecular markers for adipocytes (Cristancho and
Lazar, 2011; Tansey et al., 2004). The differentiation of a mesenchymal stem cell towards an
adipogenic cell fate as well as the intermediate steps to mature adipocyte are not well defined. A
pre-adipocyte, or adipoblast is believed to precede mature adipocytes and currently, the criteria
for identification is cell adhesion expression patterns with CD34+/CD31- from human
stromovascular fractions via cell sorting (Sengenes et al., 2005). Interestingly, when MMP
activity is completely inhibited committed adipoblasts are incapable to differentiate thus
impairing adipose tissue development in vivo (Chavey et al., 2003; Croissandeau et al., 2002;
Lijnen et al., 2002; Maquoi et al., 2002). Furthermore, MMP14 knockout mice have reductions
in adipose tissue and adipoblasts are unable to differentiate in 3D collagen gels unless the
collagen concentration levels are reduced (Chun et al., 2006).
1.1.2 Craniofacial skeletal bone diversity in zebrafish
Zebrafish are teleost fish that contain both cellular and acellular bones in the skull that can be
further classified as spongy, compact, and tubular (Weigele et al., 2016). These bone categories
can be associated with the type of ossification that occurred during craniofacial development.
Within the hypurals of the caudal fin, adipocytes have been observed similar to what I observe in
the remodeled ceratohyal bone. This is not the only location of adipocytes enclosed within bone
structures in the zebrafish. Other locations include the pectoral girdle, the anguloarticular, the
pterosphenoid, and the lower quadrate bone (Weigele et al., 2016). Interestingly, Weigele et al.
(2016) identify two distinct groups of osteoblasts, type I and type II, that associate with
perichondrium and periosteum layers in premaxilla and hyomandibular in craniofacial skeleton
respectively.
1.1.3 Periosteum and perichondrium roles during bone development
The perichondrium is first established independently from chondrogenesis of the membranous
skeleton and in long bones the perichondrium gives rise to all the populations of osteoblasts
within the periosteum and the periosteum itself (Colnot et al., 2004; Eames et al., 2003; Hall and
Miyake, 2000). Both the periosteum and perichondrium utilize Syndecan3 and Tenascin-C to
13
form the boundaries of developing cartilages and bone as well as the boundaries of condensation
size (Koyama et al., 1995; 1996). The perichondrium and periosteum serve as a boundary
distinction between chondrogenic and osteogenic cell populations apart from non-skeletogenic
mesenchyme during development. In zebrafish perichondral bones in the skull use Hedgehog
signaling via patched1 and patched2 receptors to control osteoblast differentiation in
perichondral cells (Felber et al., 2011). Twenty unique molecular distinctions between
periosteum and perichondrium in both cell layer and expression domains were defined in the
chick tibiotarsi at relevant skeletogenic time points like the onset of osteogenesis and during
chondrogenesis (Bandyopadhyay et al., 2008). Perichondrium within in vitro studies have
demonstrated that perichondrium is not required for chondrogenesis to proceed but interacts with
adjacent chondrocytes during differentiation (Liu et al., 2002). Perichondrium works in a
feedback loop with prehypertrophic chondrocytes via Ihh and PTHrP to regulate chondrocyte
differentiation (Lanske et al., 1996; Vortkamp et al., 1998). Further, Fgf18 signaling from the
perichondrium promotes proliferation during chondro- and osteogenesis in mice utilizing Runx2
(Liu et al., 2002; Kobayashi et al., 2002; Hinoi et al., 2006). The periosteum is primarily
identified as a two layered region around the surface of bone, the outer fibrous layer contains
fibroblasts and blood vessels in addition to collagens and elastin (Allen et al., 2004). The inner
cambium layer is composed of undifferentiated skeletal progenitor cells and osteoblasts (Malizos
and Papatheodorou, 2005). Bandyopadhyay et al. (2008) identified Decorin, Col1A2, and 1-76 as
distinct molecular markers that rapidly increased expression in periosteal cells and the initial
osteoblasts during the bone collar formation in the chick tibiotarsus.
1.1.4 Bone remodeling during growth
The vertebrate skeleton, including teleosts, on the whole is an organ system that regularly uses
cell fate plasticity for maintenance and growth despite the appearance of a static organ system
from a gross anatomical level (Witten and Hall, 2015). In zebrafish the mechanical load affects
ossification of the hypurals in the tail. In work performed by Van der Meulen et al. (2005) and
Fiaz et al. (2012) endurance swim training can accelerate the rate of ossification by a 15 day
reduction. These bone functional adaptations were not limited to the fin and tail but also
observed in skeletal structures in the head. In mammalian long bones the remodeling of the bone
is the coordinated actions by osteoblasts and osteoclasts generating and absorbing bone matrix.
In addition, to bone matrix modifications the bone remodeling during growth also is closely tied
14
to vascularization (Gerber et al., 1999; Eshkar-Oren et al., 2009). Throughout the eventual
marrow cavity, the chondrocyte template that was initially generated in endochondral bone
growth is progressively replaced. The trabecular bone is formed by utilizing the scaffolding
provided by calcified hypertrophic chondrocytes of the growth plate (Frisén, 2002). Trabeculae,
or bone spicules, are arranged and observable with increased density moving towards the growth
plates within the lining of the bone marrow cavity (Erlebacher et al., 1995).
1.2 Cell fate plasticity
Cell differentiation is classically depicted as a hierarchy of branching cell groups, the further a
cell sits up the hierarchy the greater the potential to generate specialized cell types observed in
the organism. The term stem cell is a reflection of this analogy, a cell that all other cells stem
from, and developmental researchers have refined the tiers of individual tissues to reflect the
diversity of fully differentiated cell types. Another ubiquitous piece of imagery when discussing
cell fate differentiation decisions is Waddington’s landscape as a central concept of
developmental biology, especially when thinking about the epigenetic landscape of a cell.
However, the caveat of a marble rolling downhill into branching valleys does not necessarily
include accommodation for the plasticity of cell identity. Cell fate plasticity refers to the variety
of methods that a cell identity can be changed from one specialized cell type to another.
Transdifferentiation represents a direct change from one defined specialized cell type to a
different unassociated specialized cell type. This change in cell type is rare but has been
documented in smooth muscle cells to skeletal myocytes of the esophagus (Patapoutian et al.,
1995). Dedifferentiation is the transition of a specialized cell type to a state of wider potency for
the eventual re-differentiation into an unrelated specialized cell identity. Following eye injury,
the newt regenerates the lens by the dedifferentiation of pigment epithelial cells (Eguchi, 1963).
Transdetermination is commonly associated with Drosophila imaginal discs. It represents a
progenitor cell population changing the cell fate, usually to a progenitor cell of a different
lineage commitment (Maves and Schulbiger, 2008). Yechoor et al. (2009) also presented
evidence that hepatic progenitor cells in the liver can lineage switch to pancreatic islet-like cells
expressing major islet hormones like insulin. Caveats to mislabeling cell fate plasticity are
15
important to take under consideration and include heterogenous cell populations within the
skeletal tissue, incomplete perspective on cellular marker expression, and cell fusion of hybrid
tetraploid cells can result in changes in cellular reprogramming and identity (Frisén, 2002). Each
of these considerations should be investigated for skeletal cells, skeletal stem and progenitors as
well as differentiated skeletal cells.
1.2.1 Cell fate plasticity in development and injury
As part of their intrigue to the skeletal research community, periosteal cells can generate cell
types for both chondrogenesis and osteogenesis suggestive of a bipotent osteochondro progenitor
cell population. This ability has ties back to mesenchymal condensation mechanisms like cell
adhesions NCAM (Fang and Hall, 1999). Interestingly, examples of intramembranous
ossification utilizing chondrogenic programs challenge the foundation of defining skeletal
elements based on the type of ossification process: occurring either through a cartilage
intermediate or explicitly without a cartilage phase. As Hall (2015) notes: “The very existence of
intramembranous and endochondral classes of ossification separates intramembranous
ossification from any association with cartilage or a cartilaginous phase, although at the earliest
stages of their deposition of osteoid many membrane bones do stain with Alcian blue, reflecting
the proteoglycans in the unmineralized matrix.” In adult upper jaw resection skeletal preps, I
have observed the premaxilla stained positive with Alcian blue and not Alizarin Red in the
proximal bone segment, so the question becomes if the mineralized bone matrix becomes
unmineralized to enable bone remodeling and if periosteal cells activate a chondrogenic program
for the progression of regeneration representing a dedifferentiation to a bipotent progenitor cell
state.
Secondary cartilage formation in chick membrane bones, and mouse calvarium another
intramembranous ossified bone also demonstrates a chondrogenic potential in vitro and in vivo
while expressing prechondrogenic genes Aggrecan, Col2a1, Col10 (Nah et al., 2000; Toma et al.,
1997; Åberg et al., 2005). Skeletal cell types demonstrating cell fate plasticity has been observed
from hypertrophic chondrocytes dedifferentiating to ligament cells in the mouse pubic joint
(Crelin, 1957; reviewed in Hall, 2015; Crelin and Koch, 1965; Crelin and Koch, 1967).
Chondrocyte dedifferentiation into an adipogenic cell fate was initially proposed by Egerbacher
et al. (1995). The reverse cell fate switch from adipocyte to chondroblast was demonstrated in
16
vitro with the addition of Tgfb1, insulin, and transferrin (Huang et al., 2004). Cartilage markers
like aggrecan, type II collagen, chondroitin sulfate and keratan sulfate were observed by Huang
et al. (2004). The adipocyte capacity for dedifferentiation looks to have a role in fracture repair
as it is observed that leptin knockout mice demonstrate slower the rates of repair and can be
reversed with local leptin applications (Khan et al., 2013). It is noted that leptin expression is
involved in the inflammatory response of IL-6 and IL-1 in fracture hematomas, soft callus
formation, and direct action on osteoblasts (Axelrad et al., 2007; Wang et al., 2011). Yet another
example of skeletal cell fate plasticity has been observed in the hypertrophic chondrocytes of
mouse Meckel’s cartilage by a series of publications by Ishizeki and colleagues. In vitro and in
grafted experiments, they demonstrate that Mmp1 secretion by hypertrophic chondrocytes
releases the cells from cartilage ECM and the cells switch to an osteoblast progenitor cell fate
and produce type I collagen, osteocalcin, and alkaline phosphatase (Ishizeki et al., 1999; Ishizeki,
2012). The series of studies echoes my results of hypertrophic chondrocytes through expression
of metalloproteinases, mmp9, removed from the cartilage ECM subsequently undergo a cell fate
transition to a skeletal progenitor state, instead of undergoing apoptosis, and contribute to
osteogenic cells during endochondral remodeling in zebrafish ceratohyal. Meckel’s cartilage
hypertrophic chondrocytes demonstrating an ability to cell fate switch emphasizes the need for
careful lineage analysis of the Meckel’s cartilage during zebrafish lower jawbone regeneration
and generation of the hybrid cartilage-bone callus.
1.3 Skeletal stem and progenitor cells in adults
The standard for classifying adult stem and progenitor cell populations are achieved by two
steps: a) define the contribution to specialized cell types by molecular labeling of the cells in the
living tissue of interest, then b) remove the population from the animal for in vitro culture and
introduce those cells into another animal and determine the ability of the cells to repopulate the
tissue of origin. These steps help to define cell fate potential and the capacity for self-renewal
and subsequent tissue regeneration. Cells from the bone marrow were initially identified to
operate as adult skeletal stem cells (Friedenstein et al., 1966; Tavassoli and Crosby, 1968) but
were challenging to classify with definitive genetic markers. Instead bone marrow stem cells
were refined by a combination of murine cytokine expression via FAC sorting (Morikawa et al.,
17
2009; Chan et al., 2013). Work recently used the molecular labeling of Lepr (Leptin-receptor) in
stromal cells as a main cellar source for bone generation (Zhou, et al., 2014). Other research
groups have contributed to identifying the bone marrow stem cell population with molecular and
cytokine markers like Nestin, Sca-1, PDGFR-alpha, and Grem-1 to label stem cell populations
with varied adipogenic, chondrogenic and osteogenic ability (Chan et al., 2013; Méndez-Ferrer
et al., 2010; Ambrosi et al., 2017; Morikawa et al., 2009; Worthley et al., 2015). A
comprehensive understanding of how these independently identified skeletal stem and progenitor
populations do and do not overlap has not yet been elucidated. It is likely that the heterogeneity
of adult skeletal stem cells can be defined into a hierarchy of cell fate potential much like what
has been achieved for the hematopoietic stem cell niche. One approach to achieve this objective
would be to systematically disrupt an individual population and determine their effect on the
other skeletal stem cell populations.
1.3.1 Periosteum and perichondrium roles during fracture repair
During fracture repair, the main source of skeletal progenitor cells is derived from the
periosteum with minimal contribution from distant systemic cell types. Given the plasticity
demonstrated in osteogenic, chondrogenic, and adipogenic lineage cells to produce other skeletal
cell types, a clear understanding of the contribution and extent of compensation of pre-existing
skeletal cells to respond to the fracture repair process is a key point of future research. Cellular,
molecular, mechanical and inflammatory signals throughout fracture healing are activating and
influencing the periosteal skeletal progenitor response. (Morgan et al., 2009; Einhorn et al.,
1995; Knothe Tate et al., 2008). Unstable fractures generate higher amounts of cartilage callus
while stabilized fractures produce minimal cartilage callus (Colnot et al., 2003; Lu et al., 2005).
Correspondingly, unstable fractures heal through endochondral ossification and stabilized
fractures heal through intramembranous ossification (Thompson et al., 2002). The origin of
periosteal skeletal progenitors is not defined yet the periosteum is transformed from
perichondrium during long bone development (Colnot et al., 2004).
1.3.2 Adult skeletal regeneration and repair in zebrafish
Dermal fin rays make up the caudal fin of zebrafish that regenerate following amputation and
fracture in under two weeks across adulthood (Azevedo et al., 2011; Itou et al., 2012). The
18
caudal fin is composed a small number of cell types, but the only skeletal cells are osteoblasts
(Tu and Johnson, 2011). The regenerative blastema to contribute to fin ray growth is in part
generated by osteoblast dedifferentiation into an osteogenic proliferative progenitor. Lineage
tracing experiments with pre-existing osteoblasts demonstrated that while the osteoblasts revert
to a progenitor state, the plasticity to generate other skeletal cell types was not observed (Knopf
et al., 2011; Sousa et al., 2011). Interestingly, the ablation of pre-existing osteoblasts does not
prevent regeneration of the osteoblasts, and another cell source is capable of generating
osteoblasts de novo (Singh et al., 2012). There is suggestive evidence, similar to what is known
in Urodele limb regeneration, that a fibroblast cell type may have osteogenic potential as sox9
and cola2a expression are observed following osteoblast ablation (Singh et al., 2012). Geurtzen
et al. (2014) demonstrated that osteoblast dedifferentiation and regeneration is not limited to the
appendages of certain vertebrates like fish and newts but also regenerates in similar patterns in
the dermal zebrafish calvarium. Molecular markers of osteoblast maturation as well as re-
entering the cell cycle are shared between mammalian skull derived osteoblasts (Lian and Stein,
1995).
1.4 References
Åberg, T., Rice, R., Rice, D., Thesleff, I., and Waltimo-Sirén, J. (2005). Chondrogenic Potential
of Mouse Calvarial Mesenchyme. J Histochem Cytochem. 53, 653–663.
Allen, M.R., Hock, J.M., and Burr, D.B. (2004). Periosteum: biology, regulation, and response to
osteoporosis therapies. Bone 35, 1003–1012.
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26
Chapter 2
Programmed Conversion of Hypertrophic Chondrocytes into
Osteoblasts and Marrow Adipocytes within Zebrafish Bones
2 Abstract
Much of the vertebrate skeleton develops from cartilage templates that are progressively
remodeled into bone. Lineage tracing studies in mouse suggest that chondrocytes within these
templates persist and become osteoblasts, yet the underlying mechanisms of this process and
whether chondrocytes can generate other derivatives remain unclear. I find that zebrafish
cartilages undergo extensive remodeling and vascularization during juvenile stages to generate
fat-filled bones. Growth plate chondrocytes marked by sox10 and col2a1a contribute to
osteoblasts, marrow adipocytes, and mesenchymal cells within adult bones. At the edge of the
hypertrophic zone, chondrocytes re-enter the cell cycle and express leptin receptor (lepr),
suggesting conversion into progenitors. Further, mutation of matrix metalloproteinase 9 (mmp9)
results in delayed growth plate remodeling and fewer marrow adipocytes. My data support
Mmp9-dependent growth plate remodeling and conversion of chondrocytes into osteoblasts and
marrow adipocytes as conserved features of bony vertebrates.
2.1 Introduction
Vertebrate bones develop via two largely distinct processes. Intramembranous (i.e. dermal) bone,
which makes up a large portion of the skull, arises through the direct differentiation of
mesenchymal precursors into osteoblasts and then osteocytes. In contrast, endochondral bone,
which comprises the majority of the axial and limb skeletons, arises through the progressive
remodeling of an embryonic cartilage template. On the outside of developing endochondral bone,
perichondral cells mature into periosteal progenitors that contribute to the bone collar. The
cartilage templates of endochondral bone are organized into distinct zones of chondrocytes:
resting, proliferative, pre-hypertrophic, and hypertrophic. In mammals, chondrocytes at the edge
of the developing hypertrophic zone largely disappear as the cartilage matrix is degraded, a
27
process concurrent with the invasion of blood vessels, hematopoietic cells, and progenitors for
osteoblasts and marrow adipocytes (Maes et al., 2010). This growth plate remodeling contributes
to the establishment of trabecular bone, complementing the cortical bone largely derived from
the periosteum, and the marrow cavity supports continued hematopoiesis. As with mammals,
zebrafish also have intramembranous and endochondral bones. Their endochondral bones are
hollow and filled predominantly with fat yet do not support hematopoiesis as in mammals
(Witten and Huysseune, 2009; Weigele and Franz-Odendaal, 2016). It has remained unclear,
however, whether zebrafish bone arises solely through osteoblast differentiation in the
periosteum, or also through invasion of the vasculature and conversion of growth plate cartilage
to bone as in mammals. The source of marrow adipocytes also remains unclear in either fish or
mammals.
It has long been appreciated that many hypertrophic chondrocytes undergo cell death during
endochondral ossification, with osteoblasts forming from periosteal cells brought into the bone
along with the vasculature (Maes et al., 2010). At the same time, there are numerous studies
showing that cultured chondrocytes can dedifferentiate into mesenchymal progenitors and/or
transdifferentiate into osteoblasts (Shimomura et al., 1975; Mayne et al., 1976; von der Mark and
von der Mark, 1977). Recent lineage tracing studies of hypertrophic chondrocytes, using
constitutive and inducible Col10a1-Cre- and Aggrecan-Cre-based transgenes in mice, has
revealed that such transdifferentiation may also occur in vivo, with chondrocytes making a major
contribution to osteoblasts within trabecular bone and potentially also the bone collar (Yang et
al., 2014; Zhou, X. et al., 2014; Jing et al., 2015; Park et al., 2015). A limitation of these studies
is the use of population-based labeling by Cre recombination, which cannot exclude low-level
and/or leaky labeling of other cell types. It is also unclear whether hypertrophic chondrocytes
can give rise to other cell types, such as marrow adipocytes, and whether hypertrophic
chondrocytes directly transform into osteoblasts or do so through a stem cell intermediate.
Finally, it is unknown whether the ability of chondrocytes to generate osteoblasts and other cell
types is specific to mammals or a more broadly shared feature of vertebrates.
In this study, I address the long-term fate of growth plate chondrocytes in zebrafish, as well as
potential mechanisms of their fate plasticity. I use the ceratohyal (Ch) bone of the lower face as a
model. This long bone, which is derived from cranial neural crest cells, exhibits properties in
common with the long bones of mammalian limbs, including two prominent growth plates at
28
either end and a marrow cavity (Paul et al., 2016). Here, I describe remodeling of the Ch from a
cartilage template to a fat-filled bone in juvenile stages, which coincides with extensive
vascularization. Using inducible Cre and long-lived histone-mCherry fusion proteins, driven by
regulatory regions of the chondrocyte genes sox10 and col2a1a, I reveal contribution of
chondrocytes to osteoblasts, adipocytes, and mesenchymal cells within the adult Ch. In mouse,
LepR expression marks bone marrow cells that contribute to osteoblasts and adipocytes primarily
after birth (Zhou, B. O. et al., 2014). In zebrafish, I find that growth plate chondrocytes express
lepr and re-enter the cell cycle during the late hypertrophic phase, raising the possibility that
Lepr+ skeletal stem cells may derive from growth plate chondrocytes. Further, I find that delayed
remodeling of the hypertrophic cartilage zone in zebrafish mmp9 mutants correlates with a
paucity of marrow adipocytes. Unlike in mouse where Mmp9 functions in hematopoietic cells
for timely growth plate remodeling (Vu et al., 1998), I find that Mmp9 is sufficient in neural
crest-derived chondrocytes of zebrafish for growth plate remodeling. My studies reveal that
growth plate chondrocytes generate osteocytes and adipocytes in zebrafish bones, potentially by
transitioning through a proliferative intermediate.
2.2 Results
2.2.1 Remodeling of the Ch bone in juvenile zebrafish
In order to characterize the progressive remodeling of an endochondral bone in zebrafish, Dr.
Sandeep Paul and Simone Schindler performed pentachrome staining on sections of the Ch bone
from juvenile through adult stages (Figure 2-1). The Ch bone is shaped like a flattened barbell,
and here they sectioned it to reveal the thin plane of the bone (see Figure supplement 2-1A) for a
view along the thicker perpendicular plane). Unlike the unidirectional growth plates in the mouse
limb, the two growth plates of Ch are bidirectional with a central zone of compact, proliferative
chondrocytes flanked by hypertrophic chondrocytes on either side (Paul et al., 2016). Unlike in
many other fish species, the Ch bone, as with other bones in zebrafish, also contains embedded
osteocytes (Witten and Huysseune, 2009). At 11 mm standard length (SL) (approx. 4.5 weeks
post-fertilization (wpf)), the Ch contains chondrocytes throughout its length with the exception
of a small marrow space at the anterior tip. The Ch is surrounded by a thin layer of cortical bone
29
that has been shown to derive from osteoblasts located on the outside of the cartilage template
(i.e. periosteum) (Paul et al., 2016). By 12 mm SL (approx. 5 wpf), both tips of the Ch contain
marrow spaces, and on the central sides of the growth plates we begin to observe small fissures
in the cortical bone and disruption of the hypertrophic zone. By 13 mm SL (approx. 5.5 wpf),
breaks in the cortical bone become more prominent and are accompanied by further degradation
of the cartilage matrix. At later stages (16 and 19 mm SL) (approx. 7 and 9 wpf), cortical bone
regains integrity and increases in thickness, and marrow adipocytes containing LipidTOX+ lipid
vesicles are seen throughout Ch (Figure 2-1B, C and Figure supplement 2-1B). By adulthood
(one year of age), the marrow cavity is filled with large fat cells and the growth plates appear
largely mineralized. While I focus on the Ch for this study, a number of other cartilage-derived
bones in the face and fins have been reported to have a similar structure in zebrafish, including
growth plates and prominent marrow fat (Weigele and Franz-Odendaal, 2016).
30
Figure 2-1. Time-course of Ch remodeling in juvenile zebrafish
(A) Pentachrome staining of a longitudinal section through the head of a 19 mm fish. The jaw is
toward the left (anterior) and the gills toward the right (posterior). The green stain highlights the
collagen matrix of cartilage, and the reddish-brown stain the mineralized matrix of bone. The
bilateral set of Ch bones is indicated. n = 3.
31
(B) High magnification views of the Ch at successive stages show the gradual replacement of
chondrocytes in the central shaft and at each end with adipocytes (which appear white due to loss
of lipid during processing). n = 3 for each stage.
(C) Higher magnification views of the boxed regions in (B). Cortical bone appears reddish-
brown. Note the breaks in cortical bone toward the lower part of the images at 12 and 13 mm,
which are largely resolved by 16 and 19 mm. Scale bars = 50 µM.
Figure supplement 2-1. Ch bone and marrow fat structure
(A) Dissected Ch bone from a juvenile zebrafish (16 mm SL) shows staining of cartilage cells by
col2a1a:GFP (green) and mineralized bone matrix by Alizarin Red. A superficial confocal
section shows cortical bone along the length of the Ch, and a central section shows a lack of
bone inside the Ch. Brightfield image of the same Ch shows large lipid droplets characteristic of
marrow adipocytes. GP, growth plate.
(B) Confocal projection of a dissected Ch (16 mm SL) from a col2a1a:GFP animal shows
cartilage in green and adipocytes stained with LipidTOX in red. Brightfield image of the same
Ch shows large lipid droplets overlapping with LipidTOX signal. n = 2. Scale bars =100 µm.
(C) Confocal section from a double transgenic animal (20 mm SL) shows fli1a:GFP+ blood
vessels (green), col2a1a:H2A:mCherry-2A-GFPCAAX chondrocytes (red nuclei), Calcein
32
Blue+ cortical bone (blue, outlined in white), and adipocytes (brightfield, grey). Note that cross-
sectional and longitudinal slices through the blood vessels in the central portion of the marrow
cavity (to the right of the chondrocytes and between the cortical bone). Scale bars =100 µm
(A,B), 50 µm (C).
2.2.2 Vascularization of the Ch bone in juvenile zebrafish
Given the transient breakdown of cortical bone, I examined whether this coincides with
vascularization of Ch. To do so, I performed confocal imaging of the dissected Ch from fish
carrying both a chondrocyte-specific col2a1a:mCherry-NTR transgene and fli1a:GFP (Figure 2-
2). I used fli1a:GFP to label endothelial cells of the vasculature, but I also noticed that
presumptive resting chondrocytes in the middle of the growth plate express this transgene. At 10
and 11 mm SL, vessels expressing fli1a:GFP are found largely on the outside of the Ch bone. By
13, 16, and 20 mm SL, I observe increasing numbers of capillaries within the Ch, coinciding
with the replacement of col2a1a:mCherry-NTR+ chondrocytes with adipocytes. High-
magnification confocal sections, performed by Dr. Sandeep Paul, at 18 mm SL clearly show
fli1a:GFP+ vessels in intimate association with adipocytes and within the Calcein Blue+ bone
collar (Figure supplement 2-1C). Given the more complicated expression pattern of fli1a:GFP, I
also independently confirmed blood vessel identity with kdrl:GFP. At 28 mm SL (approx. 26
wpf), the Ch is heavily supplied with both kdrl:GFP+ blood vessels and lyve1:DsRed+ lymphatic
vessels, which abut each side of the growth plate (Figure 2-2B). Hence, as in mammalian bones,
remodeling of the Ch bone is accompanied by extensive vascular invasion of the cartilage
template.
33
Figure 2-2. Vascularization of the Ch
(A) Confocal projections of dissected Ch bones at five successive stages. Merged fluorescent and
brightfield channels show the gradual replacement of the cartilage with a fat-filled core.
col2a1a:mCherry-NTR highlights chondrocytes that become increasingly restricted to two
growth plates (GP) at either end of the bone. fli1a:GFP labels endothelial cells and chondrocytes
located in the central portions of the growth plates. Vascularization of the Ch increases over
time. n = 2 at each stage.
(B) Confocal projection shows networks of kdrl:GFP+ vascular endothelial and lyve1:DsRed+
lymphatic endothelial cells within an adult Ch bone. The inset shows a single confocal section
through the boxed portion of the growth plate, with both blood and lymphatic vessels abutting
the edges but not penetrating into the growth plate. n = 2. Scale bars = 100 µm (A) and 200 µm
(B).
34
2.2.3 Contribution of sox10-lineage cells to osteoblasts, adipocytes, and
mesenchymal cells
Given the extensive remodeling and vascularization of Ch, I next investigated the long-term fate
of growth plate chondrocytes by multiple, independent methods. First, Dr. Joanna Smeeton
constructed an inducible sox10:CreERT2 line and crossed it to a ubiquitous bactin2:loxP-
tagBFP-stop-loxP-DsRed reporter (Blue to Red conversion: B>R). The zebrafish sox10 promoter
drives expression in early cranial neural crest cells from 10-16 hpf, followed by a second wave
of expression in all chondrocytes from 2 dpf onwards (Dutton et al., 2008). Here, I took
advantage of this second wave of expression to label developmental chondrocytes. Upon
addition of 4-hydroxytamoxifen (4-OHT) at 15 dpf, I observed extensive labeling of
chondrocytes within 5 days, as well as some cells in the perichondrium surrounding Ch and other
cartilages (Figure 2-3A). I did not observe leaky conversion in the absence of 4-OHT at either
embryonic or adult stages (Figure supplement 2-2A). I then converted sox10/B>R fish by 4-OHT
treatment at 14 dpf and raised these to adulthood (27 mm SL) for analysis, with inclusion of an
ocn:GFP transgene allowing us to identify co-labeled osteoblasts (Figure 2-3B). Confocal
maximum intensity projections through Ch revealed extensive labeling of growth plate
chondrocytes. I also observed numerous cells throughout the Ch, including in and around the
marrow cavity. In sections through the middle of Ch, I observed labeling of a number of large
diameter cells of adipocyte morphology (Figure 2-3C). In superficial sections through the
cortical bone, I also observed labeled cells that were positive for the osteoblast marker ocn:GFP,
as well as some labeled cells negative for ocn:GFP that may represent bone progenitors or other
cell types (Figure 2-3D). As a comparison, I used a constitutive Cre driven by a human SOX10
enhancer that drives expression throughout the neural crest lineage (note that this human
enhancer lacks the second wave of chondrocyte expression seen with the zebrafish regulatory
region used for the sox10:CreERT2 line) (Kague et al., 2012). When crossed to the B>R line, this
neural crest-specific SOX10:Cre line drives broader conversion in the 5 dpf head than the later
conversion of sox10:CreERT2 (Figure supplement 2-3A). Analysis of the adult Ch in SOX10:Cre
fish shows labeling of all growth plate cartilage, as well as most if not all adipocytes and
numerous smaller cells throughout the marrow and cortical bone surface (Figure supplement 2-
3B,C). This is consistent with previous studies showing that the Ch bone is neural crest-derived
(Schilling and Kimmel, 1994). However, I also detect unconverted cells in the marrow,
35
consistent with contribution of non-neural crest-derived cells such as the mesoderm-derived
vasculature (Figure 2-2).
As sox10:CreERT2-mediated conversion at 14 dpf also labels cells outside the cartilage, which
could also contribute to osteoblasts and adipocytes, I next examined animals with lower
conversion efficiency to follow discrete growth plate clones. When analyzed at 30 mm SL,
growth plate clones could be quite large, consistent with clonal selection as described in the
zebrafish heart and skeletal muscle (Gupta and Poss, 2012; Nguyen et al., 2017). In one example
with three discrete clones, I observed a clone that contributed to a narrow column of growth plate
chondrocytes in the middle of Ch that was contiguous with mesenchymal cells and then
adipocytes toward the central marrow cavity (Figure 2-3E). To more definitively follow growth
plate clones, I also examined sox10:CreERT2; ubb:Zebrabow fish in which 4OH-T treatment at
14 dpf resulted in conversion of RFP (red) to various color combinations of CFP, YFP, and RFP
at 23 mm SL. Analysis of uniquely colored clones in the growth plate revealed those that
contained both chondrocytes and adjacent adipocytes in the marrow, as well as those that
contained chondrocytes and cells embedded in cortical bone, consistent with an
osteoblast/osteocyte identity (Figure 2-3F). These clonal contributions are consistent with
chondrocytes giving rise to adipocytes and mesenchymal cells, and potentially osteoblasts, after
growth plate remodeling in zebrafish.
36
Figure 2-3. Contribution of sox10+ chondrocytes to osteoblasts and marrow adipocytes
(A) Confocal projection of a sox10:CreERT2; bactin2:tagBFP>DsRed animal treated at 15 dpf
with 4-OHT and imaged at 20 dpf. A ventral view of the lower face shows conversion in growth
plate (GP) Ch chondrocytes, as well as additional mesenchymal cells throughout the face. n = 3.
(B) Confocal projection of a dissected Ch bone from a sox10:CreERT2; bactin2:tagBFP>DsRed;
ocn:GFP animal converted at 14 dpf and imaged as an adult (27 mm SL). In addition to labeling
of the growth plates (GP), extensive DsRed+ cells are seen throughout the Ch in 3/3 strongly
converted animals.
(C) Higher magnification confocal section through the boxed region in (B) shows a subset of
adipocytes labeled by DsRed (red, arrowheads).
37
(D) Higher magnification confocal section through the boxed region in (B) shows a mixture of
converted (yellow, arrowheads) and unconverted (green) ocn:GFP+ osteoblasts, as well as
converted ocn:GFP- mesenchymal cells (red, arrows).
(E) Confocal projection of a dissected Ch bone from a sox10:CreERT2; bactin2:tagBFP>DsRed
animal converted at 14 dpf and imaged as an adult (30 mm SL). Three prominent clones in the
growth plate are numbered. In the boxed regions to the right, a discrete clone of growth plate
chondrocytes transitions into a stream of mesenchymal cells and then a number of adipocytes
(arrowheads). The brightfield image from the same sample (below) shows the lipid vesicles
characteristic of adipocytes. Similar clonal contributions were seen in 4 independently converted
animals.
(F) Confocal projection of a portion of a dissected Ch growth plate from a sox10:CreERT2;
Zebrabow animal converted at 14 dpf and imaged as an adult (23 mm SL). Images are shown
with and without the Nomarski channel. Unconverted cells are red, and distinctly colored growth
plate clones are visible. Magnified images corresponding to the boxed regions are shown without
the red channel to highlight distinct green and teal clones (brackets). The teal clone of growth
plate chondrocytes is contiguous with two similarly colored adipocytes (Ad1, Ad2), and the
green clone is contiguous with faintly green cells (arrowheads) in cortical bone. In the adipocyte
clone, the arrow indicates a green marrow cell distinct from the teal-colored adipocytes.
Comparable clonal contributions were seen in 3 independently converted animals. Scale bars =
100 µm (A), 200 µm (B,E,F), 50 µm (C).
38
Figure supplement 2-2. Characterization of the sox10:CreERT2 and col2a1a:CreERT2
transgenic lines
(A) Confocal projections of sox10:CreERT2; bactin2:loxP-tagBFP-stop-loxP-DsRed animals
treated with or without 4-OHT at 5 dpf. At 12 dpf, labeling (red) is seen in Ch chondrocytes and
some additional cells in the face. In the adult dissected Ch (31-32 mm SL), labeling is seen in the
growth plate (GP) and throughout the bone. No labeling is seen in the absence of 4-OHT. n = 4
for each treatment.
(B) Confocal projections of col2a1a:CreERT2; bactin2:loxP-tagBFP-stop-loxP-DsRed animals
treated with or without 4-OHT at 5 dpf and then re-imaged. At 16 dpf, labeling (red) is seen only
39
in chondrocytes (shown here for the Ch cartilages). In the adult dissected Ch (27-30 mm SL),
labeling is seen in growth plate cartilage and throughout the Ch bone. No labeling is seen in the
absence of 4-OHT. n = 4 for each treatment. Scale bars = 50 µm (12-16 dpf), 400 µm (adults).
Figure supplement 2-3. Neural crest contributions to the Ch bone and marrow adipocytes
(A) In SOX10:Cre; bactin2:loxP-tagBFP-stop-loxP-DsRed animals, the human SOX10 promoter
drives Cre expression and subsequent recombination of tagBFP (blue) to DsRed (red) in neural
crest lineage cells (as opposed to the zebrafish sox10 promoter of Figure 2-3, which has
additional later expression throughout chondrocytes of both crest and mesoderm origin).
Confocal projection of a zebrafish embryo (5 dpf) shows nearly all of the chondrocytes (ch) and
perichondral cells (pc) are labeled red. n = 4.
(B) Confocal projection of a dissected Ch bone from an adult zebrafish (27 mm SL). Nearly all
growth plate (GP) chondrocytes are labeled red and numerous red lineage cells are seen
throughout the marrow cavity and along the cortical surface. n = 4.
(C) Merged and separate channels corresponding to the boxed region in (B), which is centered
on the growth plate. The brightfield channel shows the many lipid droplets characteristic of
adipocytes. The red channel shows the circular profile of the cytoplasm surrounding the lipid
droplets in labeled adipocytes, as well as numerous smaller mesenchymal cells, some of which
40
are likely osteoblasts on the cortical surface. The blue channel shows the presence of cells of
non-neural-crest-origin, such as endothelial cells of mesoderm origin that form the vasculature
within Ch (see Figure 2-2). Similar contributions were observed in 4 independent animals. Scale
bars = 50 µm (A), 200 µm (B,C).
2.2.4 Contribution of col2a1a+ chondrocytes to osteoblasts, adipocytes,
and mesenchymal cells
As sox10-CreERT2-mediated conversion was broader than just the cartilage, Dr. Sandeep Paul
also generated an inducible col2a1a-CreERT2 line to more precisely trace chondrocytes and
their derivatives. In mice, Col2a1:CreERT2-mediated conversion at embryonic and early
postnatal stages broadly labels not only chondrocytes but also osteochondro progenitors, such as
those in the perichondrium and periosteum (Ono et al., 2014). In zebrafish, col2a1a is similarly
expressed at high levels in chondrocytes and in weaker levels in osteoblasts and perichondrium,
although direct evidence for col2a1a marking osteochondro progenitors in zebrafish is lacking
(Eames et al., 2012). In order to restrict expression to chondrocytes, thus avoiding potential
complications of labeling osteoblasts and putative perichondral progenitors, Dr. Paul utilized a
chondrocyte-specific “R2” enhancer of the zebrafish col2a1a gene that we had previously
characterized (Dale and Topczewski, 2011; Askary et al., 2015). Treatment of col2a1a/B>R fish
with a single dose of 4-OHT at 5 dpf resulted in extensive labeling of col2a1a-BAC:GFP+
chondrocytes at 12 dpf, but no labeling of the perichondrium, periosteum, and osteoblasts as
marked by the sp7:GFP transgene (Figure 2-4A,B; Figure supplement 2-2B). I also observed
labeling of the notochord in larval fish, but no labeling of the vasculature, blood, or other tissues
examined. After conversion of col2a1a/B>R chondrocytes at 5 dpf and examination at adulthood
(30 mm SL), maximal intensity projections through Ch revealed extensive labeling of growth
plate chondrocytes, as well as cells throughout the bone and marrow cavity (Figure 2-4C). Thus,
the majority of adult growth plate chondrocytes in Ch appear to derive from embryonic
chondrocytes. Moreover, optical sections revealed cytoplasmic DsRed staining in lipid-filled
adipocytes, presumptive osteoblasts lining the inner surface of bone, and mesenchymal cells
within the marrow cavity. In some animals displaying lower conversion efficiency, I observed
apparent clones of cells containing growth plate chondrocytes, large adipocytes, and osteoblasts
embedded in Calcein+ mineralized matrix (Figure 2-4D). Analysis of individual sections at
higher magnification revealed contribution of col2a1a-lineage cells to a subset of osteoblasts
41
within both the endosteal and periosteal surfaces of bone, as well as embedded osteocytes with
characteristic cellular processes (Figure 2-4E-G).
Figure 2-4. Contribution of col2a1a+ chondrocytes to osteoblasts and marrow adipocytes
(A, B) Ventral views of col2a1a:CreERT2; bactin2:tagBFP>DsRed animals treated at 5 dpf with
4-OHT and imaged at 12 dpf. Confocal sections and projections as indicated demonstrate
specific conversion (red) throughout cartilage, as shown by co-localization with the chondrocyte-
specific marker col2a1a-BAC:GFP (A) and lack of co-localization with the osteoblast and
periosteum marker sp7:GFP (B). Boxed areas are magnified in the top right insets. n = 6 for
each.
(C) After conversion of col2a1a:CreERT2; bactin2:tagBFP>DsRed animals at 5 dpf, a confocal
projection through the dissected Ch of an adult (30 mm SL) shows extensive DsRed+ cells in the
growth plates (GP) and throughout the bone. A higher magnification view of the boxed region,
42
along with brightfield, shows DsRed fluorescence in the thin cytoplasm surrounding the
prominent lipid vesicles indicative of marrow adipocytes, as well as in osteoblasts (osteo) of
cortical bone and mesenchymal cells (mes) within the marrow cavity. The dashed line in the x-y
slice shows the position of the x-z slice above. n = 10.
(D) In this example of a col2a1a:CreERT2; bactin2:tagBFP>DsRed animal converted at 5 dpf
and imaged as an adult, a prominent clone of DsRed+ cells are evident at the bottom of the
growth plate, consisting of GP chondrocytes, adipocytes (Ac), and osteoblasts (Ob) associated
with Calcein Green+ cortical bone. Similar clonal contributions were seen in 4 independently
converted animals.
(E) Pentachrome staining of a portion of the Ch growth plate at 19 mm SL shows the endosteum
and periosteum. Note that zebrafish have osteocytes embedded in their cortical bone (the dark
nuclei in the reddish-brown matrix).
(F, G) High-magnification images of a section of Ch cortical bone from an animal converted at 5
dpf and imaged at 28 mm SL. DsRed+ osteoblasts/osteocytes are seen in the endosteal surface
(arrows), periosteal surface (arrowheads), and embedded in bone (double arrowhead). The
merged brightfield and fluorescence image from a different example (G) shows a DsRed+ cell
with cellular processes characteristic of osteocytes. Scale bars = 100 µm (A,B), 200 µm (C,D),
50 µm (F), 20 µm (G).
2.2.5 Long-lived Histone2A-mCherry protein reveals contribution of
col2a1a+ chondrocytes to osteoblasts
A limitation of CreER-mediated lineage tracing is that it can be difficult to rule out contributions
from rare converted cells outside the population of interest, for example in the perichondrium
and periosteum. I therefore employed a Cre-independent approach to independently assess the
fate of growth plate chondrocytes. To do so, Dr. Sandeep Paul expressed a Histone2A-mCherry
fusion protein in col2a1a+ cells. An advantage of this type of lineage approach is that the levels
of Histone2A-mCherry protein, which is stably incorporated into chromatin, reflect those of
endogenous col2a1a in chondrocytes provided continued cell division does not dilute out the
fusion protein. This is in contrast to Cre-mediated approaches in which a strong ubiquitous
promoter determines the level of a reporter protein; hence, even low levels of Cre recombinase
activity outside of chondrocytes (e.g. in osteochondro progenitors) can result in strong reporter
expression. In zebrafish, col2a1a is expressed at high levels in the proliferative zone of the Ch
43
growth plate and then downregulated in the hypertrophic zone (Paul et al., 2016). In a newly
generated col2a1a:Histone2A-mCherry-T2A-GFP-CAAX transgenic line, membrane-localized
GFP-CAAX, which is rapidly turned over, is seen primarily in the proliferative zone, whereas
Histone2A-mCherry is seen uniformly throughout the Ch cartilage, confirming the long-lived
nature of this fusion protein (Figure 2-5A and Figure supplement 2-4A,B). At 7-8 mm SL
(approx. 3 wpf), which is well before the start of growth plate remodeling at 11-12 mm SL, all
Ch chondrocytes are Histone2A-mCherry+ and I do not detect Histone2A-mCherry+ cells
associated with Calcein Blue+ bone or co-expressing the osteoblast transgene ocn:GFP (Figure
2-5A and Figure supplement 2-4C,D). In contrast, at post-remodeling stages (12 and 18 mm SL),
I observe extensive overlap of Histone2A-mCherry with ocn:GFP+ osteoblasts associated with
cortical bone (Figure 2-5B and Figure supplement 2-4E). We also observe numerous Histone2A-
mCherry+ cells embedded in the endosteal and periosteal surfaces of the Ch bone (labeled by
Calcein Blue), with several of these cells co-expressing the pre-osteoblast transgene
RUNX2:GFP or the early osteoblast transgene sp7:GFP (Figure 2-5C-E). Note that ocn:GFP,
RUNX2:GFP, and sp7:GFP can all be readily distinguished from membrane GFP-CAAX by their
much stronger and cytoplasmic expression. In addition, I observed that sp7:GFP+ osteoblasts
further from the growth plate tended to have weaker Histone2A-mCherry signal than those more
closely associated with the edge of the hypertrophic zone, suggesting that hypertrophic
chondrocytes and/or their osteoblast derivatives undergo cell division to dilute out the
Histone2A-mCherry signal. These findings independently confirm the conclusions of our CreER
lineage tracing studies that col2a1a+ chondrocytes generate osteoblasts in zebrafish.
44
Figure 2-5. Tracing of col2a1a-lineage cells by a long-lived Histone2A-mCherry fusion
protein
(A) At a stage preceding growth plate remodeling (8 mm SL), col2a1a:H2A-mCherry-2A-
GFPCAAX labels chondrocytes, but not osteoblasts associated with Calcein Blue+ mineralized
bone. Growth plate (GP) chondrocytes co-express the nuclear Histone2A-mCherry protein (red)
and the membrane-localized GFPCAAX protein (green). In the middle and poles of Ch,
hypertrophic chondrocytes retain the long-lived H2A-mCherry protein but not the short-lived
GFPCAAX protein, reflective of the down-regulation of col2a1a expression during hypertrophic
maturation. n = 3.
(B) In a confocal section through the dissected Ch of a juvenile fish (18 mm SL), numerous
H2A-mCherry+; ocn:GFP+ cells are seen in regions where the cartilage template is being
converted to fat. Magnification of the boxed region shows the brightfield image (white), a
45
merged image of H2A-mCherry+ cells (red) and ocn:GFP+ cells (green), and individual channels
below. I observed a number of H2A-mCherry+; ocn:GFP+ cells (arrowheads) in 4/4 animals.
(C) Confocal projection of a dissected Ch at 18 mm SL reveals cells expressing nuclear
Histone2A-mCherry (red) on both the endosteal surface (arrows) and periosteal surface
(arrowheads) of Calcein Blue+ cortical bone. Some H2A-mCherry+ cells associated with bone
also co-express the osteoprogenitor marker RUNX2:GFP (yellow arrow). Note that the
membrane GFPCAAX signal from the col2a1a:H2A-mCherry-2A-GFPCAAX transgene is
much weaker and barely detectable in the proliferative zone at the gain settings used to image
cytoplasmic RUNX2:GFP. n = 3.
(D) Confocal section through the Ch at higher magnification shows several H2A-mCherry+;
RUNX2:GFP+ cells (yellow arrows) in the marrow cavity and close to the endosteal surface of
the Calcein Blue+ bone.
(E) In adult fish (26 mm SL), several H2A-mCherry+ cells are found to co-express the osteoblast
marker sp7:GFP on the endosteal surface. H2A-mCherry tends to be stronger closer to the
growth plate (GP); arrowheads denote stronger and arrows denote weaker H2A-mCherry signal.
The white dotted line in the x-y section shows the location of the x-z section above. n = 5. Scale
bars = 50 µm (A,D,E), 100 µm (B,C).
46
Figure supplement 2-4. Characterization of the col2a1a:Histone2A-mCherry-2A-
GFPCAAX line
(A) Confocal projection from a ventral view shows that the majority of chondrocytes co-express
nuclear H2A-mCherry (red) and membrane GFPCAAX (green) in the facial cartilages of
col2a1a:H2A-mCherry-2A-GFPCAAX fish at 7 dpf. Note the lack of fluorescent signal outside
47
cartilage, demonstrating specificity. M, lower jaw Meckel’s cartilage; Pq, upper jaw
palatoquadrate cartilage; Ch, ceratohyal cartilage. n = 10.
(B) Magnified confocal section of a region of the M cartilage from (A) shows membrane
localization of GFPCAAX (green) and nuclear localization of H2A-mCherry (red). The arrow
depicts a chondrocyte in anaphase.
(C) Confocal projection shows the bilateral set of Ch cartilages in col2a1a:H2A-mCherry-2A-
GFPCAAX; ocn:GFP fish at 7 mm SL. Merged and individual channels show that H2A-mCherry
labels chondrocytes but does not co-localize with ocn:GFP+ osteoblasts. The much weaker
GFPCAAX signal is not apparent at the lower gain used to image ocn:GFP expression. n = 2.
Scale bars = 100 µm.
(D) Dissected Ch cartilage from (C) shows lack of H2A-mCherry expression in ocn:GFP+
osteoblasts in the bone collar at this stage (prior to growth plate remodeling). The boxed region
is magnified below. Note the nuclear fragmentation (arrowheads) in several hypertrophic
chondrocytes at this stage, suggesting that some hypertrophic chondrocytes may already be
initiating a cell death program. n = 2.
(E) Confocal section of a dissected Ch cartilage from a col2a1a:H2A-mCherry-2A-GFPCAAX;
ocn:GFP fish at the beginning of growth plate remodeling (12 mm SL). Magnification of the
boxed region (1) shows a merged image of H2A-mCherry+ cells (red), ocn:GFP+ cells (green),
and the DIC channel (white). Note several lipid-filled adipocytes (arrowheads) within the distal
end of the Ch bone. Removal of the DIC channel (1’) reveals several H2A-mCherry+; ocn:GFP+
cells (arrows), with a corresponding digital section through the X-Z plane (3) revealing that these
double-positive cells reside on the bone surface and thus likely represent osteoblasts. In contrast,
an X-Z digital section through the central shaft of the Ch bone (2) shows no double-positive cells
on the cortical surface, consistent with this portion of the growth plate having not yet initiated
remodeling and hence lacking chondrocyte-derived osteoblasts. n = 2. Scale bars = 50 µm
(A,C,E), 20 µm (B,D).
2.2.6 Hypertrophic chondrocytes re-enter the cell cycle and express lepr
The conversion of hypertrophic chondrocytes to osteoblasts and adipocytes could occur in the
absence of cell division (i.e. “transdifferentiation”) and/or through partial dedifferentiation into a
proliferative progenitor. To test these possibilities, I first used incorporation of
48
bromodeoxyuridine (BrdU) to detect proliferative cells in the Ch during remodeling stages
(Figure 2-6A,B). At the beginning of remodeling (11 mm SL), I detected BrdU+ cells in the
central zone of chondroblasts in the growth plate, as well as in the perichondrium and
periosteum. In addition, I observed BrdU+ cells at the edge of the hypertrophic zone where the
cartilage matrix is being actively degraded, similar to what has been reported in mouse (Park et
al., 2015). I observed BrdU incorporation in similarly positioned hypertrophic chondrocytes at
15 and 19 mm SL, with BrdU+ cells becoming fewer in the perichondrium and periosteum by 19
mm.
I next tested whether hypertrophic chondrocytes that re-enter the cell cycle also express known
skeletal stem cell markers. In mice, LepR expression marks a heterogeneous population of cells
in endochondral bone, including a putative postnatal skeletal stem cell population (Zhou, B. O. et
al., 2014). First, based on previously unpublished work of Dr. D’Juan Farmer and Claire Arata,
we examined expression of lepr and found dynamic expression in the zebrafish Ch endochondral
bone from juvenile through adult stages. A comparison with the hypertrophic chondrocyte and
early osteoblast marker runx2b shows higher lepr expression in proliferative versus hypertrophic
chondrocytes at 8, 12, and 20 mm SL stages, with chondrocyte expression decreasing by 27 mm
SL (Figure supplement 2-5A). During remodeling of zebrafish Ch (15 mm SL), I also observe
lepr+ cells in the marrow cavity that are derived from chondrocytes, based on labeling by
col2a1a/B>R (Figure 2-6C), and I continue to observe lepr+ cells in the Ch marrow at later
stages (20 and 27 mm SL) (Figure supplement 2-5A). lepr expression appears to be modulated
progressively from enrichment in proliferative chondrocyte zone to less enrichment in
hypertrophic chondrocytes (Figure 2-6D). By comparison, Dr. Farmer observed mouse LepR
expression in the mouse femur (8 weeks old) enriched in columnar and perichondral
chondrocytes but not enriched in hypertrophic chondrocytes (Figure supplement 2-5B). These
results are consistent with lepr+ cells in the bone marrow deriving from growth plate
chondrocytes in zebrafish, although direct evidence will be needed to determine if any of these
chondrocyte-derived lepr+ marrow cells behave as skeletal stem cells in zebrafish.
49
Figure 2-6. Late-stage hypertrophic chondrocytes re-enter the cell cycle and express lepr
(A) Pentachrome staining of a section through a Ch growth plate at 14 mm SL. Arrowheads
denote two examples of hypertrophic chondrocytes at the edges of the growth plates that lack
collagen-rich matrix (green) and appear to be exiting their lacunae.
(B) BrdU incorporation (pink) relative to all nuclei (Hoechst, blue) shows recently divided cells.
Fluorescent images with or without brightfield are shown for 11 and 15 mm SL stages, and
fluorescent channel only for 19 mm SL. In addition to BrdU+ cells in the proliferative zones of
the growth plates (brackets) and perichondrium (arrows), a subset of hypertrophic chondrocytes
50
at the edges of the growth plates (arrowheads) are BrdU+ at each stage. Proliferative
hypertrophic chondrocytes were seen in sections from 3 independent animals at each stage.
(C-D) Fluorescent RNAscope in situ hybridization for lepr (green) and the hypertrophic
chondrocyte and osteoblast precursor marker runx2b (white). Red signal indicates cells derived
from col2a1a/B>R chondrocytes that were converted by addition of 4-OHT at 5 dpf (detected by
anti-DsRed antibody), and all nuclei are shown in blue (Hoechst). In a section of a Ch growth
plate at 15 mm SL, the merged channel above and red/green and red/white channels below show
expression of lepr and runx2b in chondrocytes and their derivatives. In the higher magnification
view of the boxed region (D), lepr is expressed in proliferative chondrocytes, runx2b is
expressed at high levels and lepr at lower levels in early hypertrophic chondrocytes, and lepr and
runx2b are co-expressed in late hypertrophic chondrocytes and adjacent mesenchymal cells that
have been released from the growth plate. Similar expression of lepr and runx2b was seen in
sections from 4/4 independent animals. Scale bars = 50 µm (B,C), 20 µm (D).
Figure supplement 2-5. Expression of Lepr/lepr mRNA in zebrafish and mouse
endochondral bone
(A) Sections through the Ch bone were processed for RNAscope in situ hybridization using
probes against lepr (blue) and the hypertrophic chondrocyte and osteoblast marker runx2b (red).
51
Images with DIC are shown above. Arrows indicate lepr+ cells in the marrow. Magnifications
corresponding to the boxed regions show runx2b+ hypertrophic zones flanking a central lepr+
proliferative zone within the growth plate (bidirectional arrows indicate zones). By 27 mm, a
distinct lepr+ proliferative zone is no longer apparent. n = 3 for each stage.
(B) Sections through the femur in 8-week old mice were processed for RNAscope in situ
hybridization using a LepR probe (magenta), and counterstained with Hoechst to label nuclei
(blue). Magnified regions of the growth plate show broader expression in columnar chondrocytes
and only rare expression in the hypertrophic zone. Strong LepR expression is also seen within
and near the perichondrium. Comparable LepR expression was observed in sections from three
independent femurs. Scale bars = 50 µm.
2.2.7 Requirement of mmp9 for growth plate remodeling and marrow
adipocyte formation
In mice, Mmp9 has been reported to function in hematopoietic lineage cells for growth plate
remodeling, as bone marrow transplants can rescue the delay in growth plate remodeling seen in
Mmp9 mutants (Vu et al., 1998). Here, Dr. Sandeep Paul and Simone Schindler tested whether
mmp9 might have a conserved requirement for growth plate remodeling in zebrafish, including
the generation of marrow adipocytes. At the beginning of remodeling (11 mm SL), we observe
expression of mmp9 at the edge of the hypertrophic zone, with this restricted expression in late-
stage hypertrophic chondrocytes continuing through 17 mm SL stages (Figure 2-7A). Higher
magnification views show that mmp9 expression is prominent in hypertrophic chondrocytes that
appear to be exiting their lacunae. Next, Punam Patel used CRISPR/Cas9 mutagenesis to create
an early frame-shift mutation in the mmp9 gene that is predicted to abolish most if not all protein
function (Figure 2-7B). mmp9 homozygous mutants are adult viable and do not display obvious
larval craniofacial defects. Whereas trichrome staining revealed no significant differences in the
mutant Ch growth plates at 17 mm SL, by 21 mm I observed that the hypertrophic zone was
significantly larger compared to the proliferative zone in mmp9 mutants versus controls,
indicative of a delay in growth plate remodeling (Figure 2-7C,D). The defect in growth plate
remodeling was still evident at 27 mm SL, with mutants displaying a wider Ch growth plate
(Figure 2-7E,G). Strikingly, mmp9 mutants also had fewer adipocytes in the central marrow
cavity compared to stage-matched controls (Figure 2-7E,G), consistent with adipocytes deriving
from hypertrophic chondrocytes that are released from the cartilage matrix by Mmp9 activity.
52
Given mmp9 expression in late-stage hypertrophic chondrocytes, I next tested whether Mmp9
might function in chondrocytes as opposed to hematopoietic lineage cells for growth plate
remodeling and marrow adipocyte generation in zebrafish. As the Ch bone is generated from
neural crest cells, Dr. Gage Crump and I used transplantation of ubiquitously labeled wild-type
ectodermal cells into the neural crest precursor domain of unlabeled mmp9-/- shield-stage hosts
to generate wild-type Ch bones in otherwise mutant hosts. At adult stages, we were able to
recover 8 mutant recipients with contribution of wild-type cells to chondrocytes of the Ch growth
plate. In these animals, I observed a rescue of growth plate width, with a trend toward better
rescue in wild-type versus mutant regions of the growth plate (p = 0.06), as well as a trend
toward rescue of adipocyte number (p = 0.06) (Figure 2-7E-G). As a control, transplantation of
wild-type neural crest cells into wild-type animals had no effect on Ch growth plate width and
adipocyte number. These results indicate that mmp9 is required in the neural crest lineage, and
potentially chondrocytes themselves, for efficient remodeling of the growth plate and the
generation of marrow adipocytes from chondrocytes.
53
Figure 2-7. Tissue-autonomous requirement for mmp9 in cartilage remodeling
(A) Colorimetric mRNA in situ hybridization shows expression of mmp9 (blue) in sections of the
Ch at 11 and 17 mm SL. Inset shows specific expression of mmp9 in hypertrophic chondrocytes
(arrows) at the edge of the growth plate. Nuclear fast red was used as a counterstain. n = 2 at
each stage.
(B) Schematic of the mmp9 gene locus in zebrafish. Rectangles denote exons. The site of the 8
bp deletion in the el734 allele is indicated by an arrow, with specific sequence changes shown
below. This frame-shift mutation is predicted to result in an early stop codon and loss of the
catalytic metalloproteinase domain.
(C) Trichrome staining at 21 mm SL shows enlarged growth plates in the Ch bones of mmp9
mutants. The approximate regions of the proliferative zones used for quantification in (D) are
shown by the yellow ovals.
(D) Quantification of the ratio of the hypertrophic to the proliferative zones shows a delay in
remodeling the hypertrophic zone in mmp9 mutants at 21 but not 17 mm SL. I performed a
student’s t-test and show standard error of the mean.
54
(E,F) Dissected Ch bones at adult stages (27-31 mm SL) from wild-type, mmp9 mutant, and
wild-type and mutant hosts receiving wild-type donor neural crest transplants (blue). Ectoderm
cells from bactin2:tagBFP>DsRed donors were transplanted unilaterally into the neural crest
precursor domain of unlabeled hosts at 6 hpf. Red two-sided arrows indicate the width of the
posterior growth plates. (+) denotes sides receiving transplants, and (-) denotes contralateral
control sides. The rescued growth plate from the mmp9 mutant receiving a wild-type neural crest
transplant is shown at higher magnification in (F), with blue arrows showing a narrower wild-
type growth plate clone and magenta arrows a wider mutant clone.
(G) Quantification shows that mmp9 mutants have wider growth plates and fewer adipocytes in
the marrow than wild-type siblings. Wild-type neural crest transplants rescue growth plate width
in mmp9 mutants, with a trend toward better rescue in areas of the growth plate with wild-type
(blue) versus mutant (magenta) clones. There was also a strong trend toward rescue of adipocyte
number with wild-type neural crest transplants that contributed to growth plate chondrocytes. I
performed a Tukey-Kramer HSD test and show standard error of the mean. Unless indicated, all
other comparisons were not significant (p<0.05). Scale bars = 50 µm (A,C), 200 µm (E, F).
2.3 Discussion
Despite anatomical differences between zebrafish and mammalian bones, I find that growth plate
remodeling is remarkably well conserved and thus likely ancestral to bony vertebrates. The
zebrafish Ch undergoes a transient breakdown of cortical bone near the growth plates, which
coincides with extensive vascularization and an Mmp9-dependent replacement of hypertrophic
chondrocytes with fat and bone. Using multiple methods of lineage tracing, including a Cre-
independent technique, I show that late-stage hypertrophic chondrocytes generate not only
osteoblasts but also marrow adipocytes in zebrafish. Further support for the ability of
chondrocytes to generate adipocytes is that delayed growth plate remodeling in mmp9 mutants
results in a paucity of marrow adipocytes in adults. Lastly, I show that hypertrophic
chondrocytes re-enter the cell cycle and contribute to lepr+ mesenchymal cells, raising the
possibility that partial dedifferentiation into proliferative progenitors underlies chondrocyte fate
transitions inside endochondral bones.
55
As zebrafish have hollow bones that lack hematopoiesis, one possibility was that most if not all
of the cortical bone of adult zebrafish would be simply derived from the periosteum. However, I
find significant contribution of chondrocyte-derived cells to both the endosteal and periosteal
surfaces of the Ch bone, similar to what has been described in the mouse (Yang et al., 2014;
Zhou et al., 2014a; Jing et al., 2015). I also reveal chondrocytes to be a significant source of
marrow adipocytes in zebrafish. Although the incomplete conversion efficiency of the CreER
lines, as well as the expression of sox10:CreER outside of chondrocytes, made it difficult to
precisely quantify what pro-portion of osteoblasts and marrow adipocytes derive from
chondrocytes, there are likely other sources for these cells, in particular the periosteum which
houses several types of skeletal stem cells (Debnath et al., 2018). Our findings raise the question
of whether marrow adipocytes also derive in part from chondrocytes in mammals. Indeed, older
studies have demonstrated the ability of murine chondrocytes to differentiate into adipocytes in
vitro (Heermeier et al., 1994; Hegert et al., 2002). Further, Col2a1:CreER cells converted at
postnatal day three in mouse were found to give rise to adipocytes in the metaphyseal bone
marrow, although a caveat is that Col2a1:CreER marks both chondrocytes and progenitors at
early postnatal stages (Ono et al., 2014).
The finding that late-stage hypertrophic chondrocytes can re-enter the cell cycle and express lepr
suggests that at least some of these cells may dedifferentiate into stem-like cells, which
subsequently expand in number and differentiate into osteoblasts and adipocytes. Future
molecular profiling of these chondrocyte-derived marrow mesenchymal cells will be needed to
better characterize their relationship to previously identified skeletal stem cells. I also cannot rule
out that some hypertrophic chondrocytes directly change into adipocytes and/or osteoblasts in the
absence of cell division (i.e. ‘transdifferentiation’). Indeed, the detection of osteoblasts with
strong col2a1a:Histo-ne2A-mCherry signal suggests that in some cases chondrocytes can form
osteoblasts with little to no cell division, as otherwise the Histone2A-mCherry signal would have
been diluted out with successive cell divisions. On the other hand, osteoblasts farther from the
growth plate tended to have weaker Histone2A-mCherry signal, suggesting proliferation of a
progenitor intermediate, and I directly observed late-stage hypertrophic chondrocytes undergoing
cell division. Whereas it has been suggested in mouse that only chondrocytes near the periosteal
surface, that is ‘borderline chondrocytes’, may be capable of lineage plasticity (Bianco et al.,
1998; Maes et al., 2010), I detected lineage clones containing growth plate chondrocytes,
56
mesenchymal cells, and adipocytes in the central part of the growth plate (Figure 2-3E,F),
arguing against such a model.
A notable feature of LepR-lineage cells in mice is that they contribute to osteoblasts and
adipocytes primarily in postnatal phases, that is when growth plate remodeling is already
underway (Yang et al., 2014). It would therefore be interesting to test whether LepR-expressing
skeletal stem cells have a similar origin from hypertrophic chondrocytes in mammals. One
caveat is that I detect endogenous lepr expression in both marrow cells and growth plate
chondrocytes in zebrafish. However, the more specific labeling of marrow cells by the LepR-Cre
in mouse likely reflects the Cre insertion being in the long LepR isoform (containing exon 18b)
that displays more restricted expression than the short isoform (Zhou et al., 2014b). Indeed, LepR
mRNA and protein has also been reported in chondrocytes of mouse (Hoggard et al., 1997), rat,
and human (Morroni et al., 2004), a finding I confirmed in the postnatal mouse femur, including
the same higher expression in immature versus hypertrophic chondrocytes that I observe in
zebrafish (Figure supplement 2-5B). Without a comparable long-isoform lepr-Cre line in
zebrafish, I cannot therefore conclude whether zebrafish lepr+ marrow cells are comparable to
those described in mouse. The future generation of Cre lines to specifically mark lepr+ marrow
cells in zebrafish will be needed to determine whether these chondrocyte-derived marrow cells
also act as stem cells for osteoblasts and adipocytes in post-embryonic fish.
A similar delay in the remodeling of the hypertrophic zone in mouse Mmp9 and zebrafish mmp9
mutants reveals genetic conservation of growth plate remodeling from fish to mammals. In
contrast to mice, I find that Mmp9 in zebrafish appears to function primarily in chondrocytes for
growth plate remodeling. I also observed rescue of marrow adipocyte number by wild-type
chondrocytes, though this had moderate statistical significance (p = 0.06), potentially owing to
low sample size (n = 8), the mosaic contribution of wild-type cells to the growth plate, and/or
roles of cells outside the growth plate to marrow adipocyte generation. In mice, loss of Mmp13 in
chondrocytes does result in a growth plate remodeling delay, with global Mmp13 deletion
enhancing the remodeling defects of Mmp9 mutants (Inada et al., 2004; Stickens et al., 2004). It
may be that MMPs are derived from both chondrocytes and invading hematopoietic cells (e.g.
osteoclasts), with the relative importance of each cell source varying between zebrafish and
mammals. Compensation by mmp13 might also explain why I detected growth remodeling
defects in mmp9 zebrafish mutants at 21 and 27–31 mm SL stages, but not at 17 mm SL. Mmp9
57
and Mmp13 have known roles in degrading components of the cartilage extracellular matrix
(Page-McCaw et al., 2007), consistent with our observed loss of collagen-rich matrix in
hypertrophic chondrocytes at the edges of the zebrafish growth plate. Secreted Mmp9 may
therefore function simply to degrade cartilage matrix and facilitate release of dedifferentiating
chondrocytes into the marrow. Intriguingly, Mmp9 has recently been shown to have an
additional, non-canonical function in the nucleus for histone H3 tail cleavage. Whereas this has
only been demonstrated so far for osteoclasts (Kim et al., 2016), it remains possible that Mmp9
could have a similar non-canonical function in altering the chromatin structure of hypertrophic
chondrocytes to allow them to acquire new potential. In conclusion, our data support
conservation of mammalian-like growth plate remodeling in zebrafish, which provides new
opportunities for better understanding the molecular and cellular mechanisms by which
hypertrophic chondrocytes transform into osteoblasts, marrow adipocytes, and potentially adult
skeletal stem cells within endochondral bones.
2.4 Materials and Methods
2.4.1 Zebrafish transgenic lines and mmp9 mutants
All procedures were approved by the University of Southern California Institutional Animal Care
and Use Committee. Published Danio rerio lines include Tg(Has.RUNX2:EGFP)
zf259
and
Tg(Mmu.Sox10-Mmu.Fos:Cre)
zf384
(Kague et al., 2012), Tg(sp7:EGFP)
b1212
(DeLaurier et al.,
2010), Tg(Ola.Osteocalcin.1:EGFP)
hu4008
(Knopf et al., 2011), Tg(col2a1aBAC:GFP)
el483
and
Tg(col2a1aBAC:mCherry-NTR)
el559
(Askary et al., 2015), Tg(bactin2:loxP-BFP-loxP-DsRed)
sd27
(Kobayashi et al., 2014), Tg(fli1a:eGFP)
y1
(Lawson and Weinstein, 2002), Tg(kdrl:eGFP)
s843
(Jin et al., 2005), Tg(-5.2lyve1b:DsRed)
nz101
(Okuda et al., 2012), and Zebrabow -
Tg(ubb:LOX2272-LOXP-RFP-LOX2272-CFP-LOXP-YFP)
a131
(Pan et al., 2013). The
sox10:CreERT2 transgene was generated with Gateway Cloning (Invitrogen) and the Tol2 kit
(Kwan et al., 2007) by combining p5E-sox10 (Das and Crump, 2012), pME-CreERT2, p3E-pA,
and pDestTol2pA2. The col2a1a-R2-E1b:CreERT2 and col2a1a-R2-E1b:H2A.F/Z-mCherry-
P2A-GFPCAAX transgenes utilize a zebrafish col2a1a R2 enhancer element with a minimal
E1B promoter sequence (Dale and Topczewski, 2011). For col2a1a-R2-E1b:CreERT2, CreERT2
58
was amplified using pENTR/D-CreERT2 as template and primers 5’-
TTCTTGTACAAAGTGGCCACCGGCCACCATGTCCAATTTACTGACCGTACAC-3’ and
5’-TAGAGGCTCGAGAGGCCTTGTCAAGCTGTGGCAGGGAAACCCTC-3’. The amplified
PCR product was combined with an NcoI/EcoRI fragment of pDestTol2-col2a1aR2-E1B-
eGFPpA (Askary et al., 2015) using Gibson Assembly (New England Biolabs). For col2a1a-R2-
E1b:H2A.F/Z-mCherry-P2A-GFPCAAX, H2A.F/Z-mCherry was amplified using template
pME-H2A.F/Z-mCherry and primers 5’-
TTTCTTGTACAAAGTGGCCAAAGCTTGGATCCCGGCCACCATGGCAGGTGGAAAAG
CAGG-3’ and 5’-
AAGTTGGTTGCTCCCGACCCCTTGTACAGCTCGTCCATGCCGCCGGTG-3’. P2A-
GFPCAAX was synthesized as a gBlock (IDT). H2A.F/Z-mCherry and P2A-GFPCAAX were
combined with a NcoI/EcoRI fragment of pDestTol2-col2a1aR2-E1B-eGFPpA using Gibson
Assembly. All transgenes were injected at 30 ng/μL along with 50 ng/μL Tol2 mRNA into one-
cell-stage embryos, with these animals raised and outcrossed to identify stable germline
founders. CreERT2 transgenes were injected into Tg(bactin2:loxP-BFP-loxP-DsRed)
sd27
embryos, followed by overnight treatment with 10 μM (Z)-4-Hydroxytamoxifen (4-OHT)
(Sigma-Aldrich H7904) at 2 dpf and screening for DsRed+ chondrocytes at 6 dpf using a Leica
fluorescent stereomicroscope. Embryos with fluorescent cartilages were raised to adulthood and
outcrossed to identify founders. I used Tg(sox10:CreERT2)
el777
, Tg(col2a1a-R2-
E1b:CreERT2)
el712
, and Tg(col2a1a-R2-E1b:H2A.F/Z-mCherry-P2A-GFPCAAX)
el695
lines for our
experiments. Two additional alleles of Tg(col2a1a-R2-E1b:CreERT2) and one additional allele
of Tg(col2a1a-R2-E1b:H2A.F/Z-mCherry-P2A-GFPCAAX) gave similar cartilage-specific
expression. To generate mmp9
el734
I used CRISPR/Cas9 mutagenesis to target the second exon.
gRNA targeting the sequence 5’-TTGATGCCATGAAGCAGCCC-3’ was injected at 25 ng/μl
with 50 ng/μl Cas9 RNA into one-cell-stage embryos as described (Hwang et al., 2013). The
mmp9
el734
allele is an 8-bp deletion that results in the incorporation of 13 additional amino acids
after amino acid 98 (P), followed by a premature stop codon that is predicted to completely
abolish the catalytic metalloproteinase domain.
2.4.2 Histology and LipidTOX staining
Live bone staining of dissected ceratohyal bones was performed by treating with 50 µg/ml
Alizarin Red (Sigma Aldrich, cat. no. A5533), Calcein Green (Thermofisher Scientific, cat. no.
59
C481), or Calcein Blue, AM (Thermofisher Scientific, cat. no. C1429) for 5 min and repeatedly
rinsing in embryo medium as described (Paul et al., 2016). I performed adipocyte labeling of
dissected ceratohyal bones by incubating in a 1:200 solution of HCS LipidTOX Deep Red (Life
Technologies, cat. no. H34477) for 15 min and rinsing in embryo medium as described (Minchin
and Rawls, 2017). Paraffin embedding and histology were performed as described (Paul et al.,
2016). I cut blocks into 5 μm sections on a Shandon Finesse Me+ microtome (cat. no. 77500102)
and collected sections on Apex Superior Adhesive slides (Leica Microsystems, cat. no.
3800080). Pentachrome and Trichrome staining were performed according to manufacturer's
instructions (Movat-Russell modified pentachrome stain kit, Newcomer Supply cat. no. 9150A;
Gomori One-Step, Aniline Blue, trichrome stain kit, Newcomer Supply cat. no. 9176A).
2.4.3 Immunohistochemistry and in situ hybridization
For cell proliferations assays, fish were immersed in system water containing 4.5 mg/ml BrdU
(Sigma Aldrich, cat. no. B5002) bath for 1 h, followed by two washes in system water, fixation
in 4% paraformaldehyde, and paraffin embedding. Immunohistochemistry was performed as
described except that I blocked with 2% normal goat serum (Jackson ImmunoResearch, cat. no.
005-000-121). After cutting thin sections, I performed antigen retrieval by treating slides with
citrate buffer (pH 6.0) in a steamer set (IHC World, cat. no. IW-1102) for 35 min. Primary
antibodies include rat anti-BrdU (1:100, Bio-Rad, cat. no. MCA2060GA) and rabbit anti-
mCherry (1:250, Novus Biologicals, cat. no. NBP2-25157). I used AlexaFluor secondary
antibodies and Hoechst to visualize nuclei. Colorimetric in situ hybridization was performed as
described (Paul et al., 2016). The mmp9 riboprobe was generated by PCR amplification of
zebrafish genomic DNA with primers 5’-GTCTCCAATACTAAAGCTCTGAAGAAG-3’ and 5-
‘TAGGATGTCGAAGGTCTATAGAGAATG-3’ and cloning into pCR-BluntII-TOPO (Life
Technologies). RNA probe was synthesized with T7 polymerase (Roche) after linearizing the
plasmid with BamHI restriction. RNAscope probes for leptin receptor (lepr) and runx2b were
synthesized by Advanced Cell Diagnostics in Channel 1 and Channel 2, respectively, and for
mouse Lepr in channel 1. Paraformaldehyde-fixed paraffin-embedded sections were
deparaffinized, and the RNAscope fluorescent multiplex v2 assay combined with
immunofluorescence was performed according to manufacturer’s protocols and with the ACD
HybEZ Hybridization oven.
60
2.4.4 Neural crest transplantations
At 6 hpf, donor ectoderm from the animal cap of bactin2:loxP-tagBFP-loxP-DsRed embryos was
transplanted into the neural crest precursor domain of mmp9
-/-
hosts as described (Crump et al.,
2004). Embryos displaying unilateral tagBFP fluorescence in the face at 3 dpf were raised in the
nursery and then genotyped at 14 dpf for the mmp9 mutant allele. Homozygous mutant and wild-
type siblings were raised at similar density until they reached the indicated sizes for analysis.
2.4.5 Imaging
Brightfield images of pentachrome and trichrome stains, and colorimetric in situ hybridizations,
were acquired with a Zeiss AxioImager.A1 microscope and a Zeiss slide scanner AxioScan.Z1.
Focus stacking of multiple images was done in Adobe Photoshop CS5. Fluorescence images
were captured on a Zeiss LSM800 confocal microscope, with representative sections or
maximum intensity projections shown as specified. Tiling was performed using ZEN software or
manually stitched together using Fiji. Brightness and contrast were adjusted in Adobe Photoshop
CS5 with similar settings for experimental and control samples.
2.4.6 Quantification and statistical analyses
I stage-matched control and experimental zebrafish by measuring standard body length (SL)
from the tip of the snout to the edge of the hypuralia. Adipocyte counts and area/width
measurements of growth plates were calculated using Fiji. The proliferative zone of the growth
plate was defined as the central region of compact chondrocytes. All measurements were
performed blinded to genotype. Statistical significance was determined by a student’s t-test for
pair-wise comparisons or Tukey-Kramer HSD tests for multiple comparisons, using GraphPad’s
Prism 7 software.
61
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Chapter 3
Defining the Progenitor Population in Adult Zebrafish Jaw Bone
Regeneration
3 Abstract
While many bone fractures do heal on their own, fracture non-unions and more traumatic
skeletal lesions require invasive surgical interventions. As the innate capacity of bones to repair
simple fractures suggests the presence of a bone progenitor population, an attractive strategy
would be to augment the ability of these endogenous progenitors to repair larger lesions. Bone
repair often involves a cartilage intermediate, yet the source of an adult skeletal progenitor cell
type is not fully defined. I partnered with Dr. Sandeep Paul to develop a new and robust model of
bone regeneration in the zebrafish lower jaw. We find that chondrocytes co-express genes
associated with osteoblast differentiation then produce extensive bone matrix, in marked contrast
to chondrocytes during craniofacial skeletal development. The likely source of repair
chondrocytes is a population of Runx2+/Sp7− cells that emanate from the periosteum, a tissue
that contributes only osteoblasts during bone homeostasis (Figure 3-1). Therefore, a distinct
differentiation program is utilized by the periosteal subpopulation to uniquely achieve large-scale
bone regeneration. I think it is suggestive the bone-forming potential of repair chondrocytes is
due to their derivation from osteogenic cells in the periosteum.
3.1 Introduction
Approximately 8 million bone fractures occur annually and 10% of these are delayed and
nonunion fractures which fail to heal despite therapeutic intervention (Holmes et al., 2017). For
smaller breaks autograft bone is sometimes an option, but for many injuries artificial materials
must be used, which lack the biological activities of native bone (Lieberman, 2005). One
approach to this problem is to understand how lower vertebrates that naturally repair large
skeletal lesions, such as amphibians (Graver et al., 1978; Ghosh et al., 1994) and zebrafish
(Kyritsis et al., 2012; Singh et al., 2012) are able to more effectively utilize these skeletal
progenitors for repair. Large-scale bone regeneration across the vertebrates mentioned, including
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human fracture healing is the formation of a cartilage callus to connect the fracture sites,
especially in the absence of mechanical stabilization (Pritchard and Ruzicka, 1950; Lieberman
and Friedlaender, 2005).
Zebrafish is an ideal model for study due its powerful genetic tools and imaging capabilities. In
particular, the ease of generating a large number of fluorescent reporter lines that allow us to
monitor cell fate, differentiation, cell-cell interactions, and to isolate cells for genomic analysis
and ablate specific cell types. Together, these tools allow me to make novel insights into the cell
behaviors and molecular pathways underlying bone regeneration. In the future, these findings
will have the potential to lead to new ways of improving bone healing in patients.
Craniofacial bone development is highly conserved between zebrafish and humans, and hence
understanding large-scale bone regeneration in zebrafish may lead to novel treatments in
patients. Previous studies of zebrafish bone regeneration in the fin and calvaria demonstrated
bone regeneration occurs through osteoblast dedifferentiation (Knopf et al., 2011; Geurtzen et
al., 2014), or de novo osteoblast generation (Singh et al., 2012). The regeneration of the lower
jawbone through a prominent cartilage intermediate provides novel understandings into large-
scale skeletal repair in contrast to previous zebrafish skeletal regeneration studies. Further, a
unique feature of zebrafish jawbone regeneration is the involvement of an unusual bone-
producing chondrocyte population.
In mammalian endochondral bone development, chondrocytes are formed by a condensation of
Sox9+ mesenchymal cells to then produce extracellular matrix proteins including type II
collagen (Col2a1) and later, hypertrophy-specific factors like type X collagen (Col10a1) and
connective tissue growth factor (Ctgf) (Mackie et al., 2008). The hypertrophic chondrocytes
initially produce calcified matrix, but then undergo apoptosis and replacement by invading
osteoblasts (Maes et al., 2010), although increasing evidence suggests that hypertrophic
chondrocytes contribute to osteoblasts as well (Yang et al., 2014). In fact, some hypertrophic
chondrocytes escape apoptosis and generate long-lived osteocytes (Mayne et al., 1976; von der
Mark and von der Mark, 1977; Yang et al., 2014; Zhou et al., 2014). Both perichondrium and
periosteum cells contribute to bone growth and remodeling as blood vessels invade the zone of
dying hypertrophic chondrocytes (Maes et al., 2010). In the context of bone repair, it has been
widely assumed that chondrocytes also serve as a template for invading osteoblasts. However,
our data suggest a different mechanism. Instead of undergoing apoptosis, I observe that the
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chondrocytes in the regenerating mandible express many genes in common with osteoblasts and
directly make new bone.
The progenitor population that generates these hybrid chondrocytes during large-scale bone
regeneration are not well defined. During bone formation, osteoblast precursors differentiate into
immature osteoblasts with high expression levels of runx2, and subsequently sp7 (osterix) and
type I collagen (col1a1) (Ortuno et al., 2013). Osteoblast maturation includes expression of
mineralization associated genes like spp1 (osteopontin) and bglap (osteocalcin). Intriguingly,
hypertrophic chondrocytes also express the same osteoblast genes but at low levels and produce
a mineralized (i.e. calcified) matrix (Dy et al., 2012). Repair chondrocytes within the cartilage
callus of mammalian fractures produce BGLAP and bone-like collagen fibers (Bahney et al.,
2014). In the context of adult zebrafish jawbone regeneration, regenerating hybrid chondrocytes
use osteoblast-associated gene expression in different functional capacities compared to skeletal
bone development.
Following initial skeletal development, bones remain dynamic organs that remodel by osteoclasts
removing old bone and osteoblasts adding new bone in response to biomechanical forces
(Apschner et al., 2011). Bone homeostasis utilizes the periosteum, a layer of connective tissue
surrounding the bone surface, as the source of new osteoblasts (Ono et al., 2014). The
periosteum appears to contribute first to cartilage and then to bone during mammalian bone
fracture repair (Murao et al., 2013). Specifically, the Akiyama lab investigated the source of the
soft cartilage callus in mice by generating transgenic lines that label Sox9 osteochondro
progenitor cells in vivo (Murao et al., 2013). Importantly, they also identified other reporters of
multipotent mesenchymal cells that contribute to mouse limb development including Prx1, and
two distinct Col1a1 promoters, 3.6kb and 2.3kb which label premature mesenchymal cells and
mature osteoblasts respectively. Examination of the transgenic promoters in a bone fracture
system indicated that 3.6kb Col1a1 and Prx1 label both the periosteum and the cartilage soft
callus while 2.3kb Col1a1 was expressed in mature osteoblasts and osteocytes. The conclusions
found by Murao in mouse soft fracture callus is strong supportive evidence that the source of
chondrocytes expressed in the zebrafish mandible is likely generated from the periosteum.
During mammalian bone fracture healing, the inner cambium layer of the periosteum appears to
provide the progenitor cells which then produce chondrocytes and osteoblasts (reviewed in
(Mackie et al., 2008)). More recently, a periosteal associated gene, Periostin (Postn) represents
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an adult skeletal stem cell population that shows a greater regenerative potential than bone
marrow stem cells (Lageneste et al., 2018).
The zebrafish mandible also consists of the Meckel’s cartilage, an element that is present in
mammalian development during lower jaw bone formation but the chondrocytes are gradually
replaced in mice and humans (Parada and Chai, 2016). In zebrafish the Meckel’s cartilage is also
formed during skeletal development as a template for bone formation but the element is
maintained across the lifespan of the fish (Paul and Crump, 2016). Similar to the periosteum,
lining the surface of the Meckel’s cartilage is a flattened cell layer of perichondral cells
surrounding the outside of the cartilage matrix (Felber et al., 2011). In zebrafish, there are no
established definitive genetic markers of periosteal or perichondral cells.
As mentioned earlier, endochondral bone formation is one process for generating the vertebrate
skeletal system. In contrast, intramembranous ossification is an alternative skeletal formation
process that notably does not involve a cartilage based template for bone collar formation. The
Sox9+ mesenchymal stem cells directly generate osteoblasts instead of condensing to generate
chondrocyte-associated extracellular collagen matrix, specifically type I collagen (Bi, et al.,
1999; Sandberg et al., 1989). However, that does not necessarily preclude the possibility that
skeletal progenitor cells that are osteogenic utilize chondrogenic programming and early matrix
deposition to generate the intramembranous bone element.
The lower jawbone ossifies around the extensive Meckel's cartilage that differentiates early in
craniofacial development (Eberhardt et al., 2006). In contrast to the lower jawbone, the upper
jaw consists of the maxilla appearing at 4 dpf and pre-maxilla appears between 8-10 dpf as
dermal ossifying bones derived from the pharyngeal first arch (Talbot et al., 2012). Considering
the repair chondrocytes distinct expression of osteogenic genes to produce mineralized bone, the
source of repair chondrocytes likely represents an adult skeletal progenitor cell population with
hybrid bone-cartilage potential. Therefore, it is important to define the source of an adult skeletal
progenitor cell population specifically, if it is periosteum derived or generated from another
resident cell type during regeneration. Furthermore, there is value in understanding how the
periosteum, which produces only bone during homeostasis, then also produces cartilage during
repair.
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Figure 3-1. Working Model of Lower Jawbone Regeneration
(A) After resection,Runx2+/Sp7- progenitors in the periosteum produce ossifying (hybrid)
chondrocytes that contribute to regeneration of the jawbone.
(B) Runx2+/Sp7- progenitors make osteoblasts during bone homeostasis. After injury, signals
from macrophages (MΦ) may divert these progenitors to produce cartilage.
3.2 Results
3.2.1 Lower Jawbone regeneration produces a hybrid cartilage-bone
callus
In mammalian fractures, Sox9+ progenitor cells within the periosteum contribute not only to the
cartilage callus but also to cells that express the osteoblast differentiation genes Runx2 and
Sp7/Osterix (Akiyama et al., 2005; Murao et al., 2013). These studies showed that periosteal
cells that generate both bone during homeostasis and the cartilage callus during repair express
Col1a1 utilizing a periosteal enhancer located 3.6kb from its transcription start site. Similarly, in
zebrafish development, osteoblast differentiation in the periosteum proceeds through sequential
expression of runx2b, sp7/osterix, and mature osteoblast markers like col1a2 and
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bglap/osteocalcin (Li et al., 2009). Whereas zebrafish are known to utilize runx2b during fin
regeneration in adults (Singh et al., 2012; Brown et al., 2009), it is unclear if Runx2 genes are
also involved in adult bone homeostasis as observed in mammals. There is also evidence of
distinct molecular subdomains of periosteum-derived progenitors across vertebrates
(Bandyopadhyay et al., 2008), yet which subtype of progenitor gives rise to the cartilage callus
during bone repair still needs to be resolved. Shortly after injury, the periosteum appears to
generate large numbers of cartilage cells (chondrocytes), yet these cells express markers of and
appear to become long-lived bone cells (osteoblasts) (Paul et. al, 2016). Instead of undergoing
apoptosis, we observe the chondrocytes in the regenerating mandible directly make new bone by
further differentiation into osteoblasts. This observation is further supported by the endogenous
expression of the chondrocyte and osteoblast genes, col1a1a and col2a1a, within the same
regenerated cartilage cells (Figure 3-2). An important question then is the origin of these bone-
forming chondrocytes during repair.
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Figure 3-2. Lower jawbone regeneration produces a hybrid cartilage-bone callus
(A) A regenerative time course of bone regeneration demonstrates consistent patterns of cartilage
and bone. Approximately 1mm of the lower jawbone was surgically removed from adult fish at 0
day post resection. n = 4. At 7 dpr, cartilage (Alcian Blue) begins to fill in the lesion (white
arrow). n = 5. This cartilage callus increases to join with the corresponding end at 14 dpr and
regenerated bone (Alizarin Red, black arrow) progressively replaces cartilage by 30 dpr. 14 dpr:
n = 5. 30 dpr n = 5.
(B-D) Previously published in Paul et al., 2016. At 10 dpr, the fluorescent in situs of sections
of the regenerating jaw show co-expression of col2a1a with col1a1a at each end of the resected
bone as well as in the cartilage callus. High magnification regions display the cellular overlap of
co-expression. runx2b within the cartilage callus is observed.
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(E, F) col2a1a:GFP+ chondrocytes are observed surrounded by mineralized bone at 12 dpr
(Alizarin Red) and further begin to express the mature osteocyte marker bglap/osteocalcin at
30dpr.
3.2.2 Runx2+/Sp7- periosteal subpopulation gives rise to the cartilage
callus during jawbone regeneration
Given the known role of the periosteum in mammalian fracture repair, we examined a potential
role of the zebrafish jawbone periosteum in generating the cartilage callus during bone
regeneration. Histological analysis showed that the periosteum undergoes marked thickening and
caps the cut bone by 2 dpr, with increased expansion of periosteal mesenchyme by 4-6 dpr
(Figure 3-3). We also detected col1a1a expression in the periosteum of the uninjured jaw, and
proliferating BrdU+ col1a1a-expressing cells emanating from the injured periosteum at 4 dpr
(data not shown). To better understand the behavior of the periosteum during jawbone repair, we
analyzed transgenic lines that mark either pre-osteoblasts (RUNX2:GFP) or early osteoblasts
(sp7:GFP) (Figure 3-3C). While sp7:GFP labels a continuous zone of osteoblasts surrounding the
uninjured bone, RUNX2:GFP labels a subset of cells in the periosteum. Interestingly, I observed
RUNX2:GFP+ cells but no sp7:GFP+ cells streaming from the periosteum into the injury site by
4 dpr. A marked expansion of the number of RUNX2:GFP+ cells can be seen as early as 2 dpr
(data not shown), and by 7 dpr RUNX2:GFP+ chondrocytes are visible within the expanding
mesenchyme filling the lesion (Figure 3-3C). Given the long-lived nature of GFP protein, my
preliminary results might be consistent with RUNX2:GFP+ cells in the periosteum proliferating
and contributing to new mesenchyme and cartilage during repair. Alternatively, RUNX2:GFP
labeling could reflect periosteal cells turning on Runx2 gene expression in response to injury.
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Figure 3-3. Activation of the periosteum during jawbone regeneration
(A, B) Previously published in Paul et al., 2016. Hematoxylin and eosin sections of adult lower
jaw resections show that inflammatory cells invade the resected area by 2 dpr but decrease by 4
dpr (arrows in A). The periosteum cell layer thickens in response to injury (white arrowheads in
B).
(C) Previously published in Paul et al., 2016.RUNX2:GFP and sp7:GFP (black arrowheads)
label the uninjured periosteum of the adult jaw, but only RUNX2:GFP+ cells are seen
contributing to the expanding mesenchyme at 4-7 dpr (white arrows) and repair chondrocytes by
7 dpr (white arrowhead).
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3.2.3 Juvenile Lower Jaw Resection
Following lower jaw resection surgeries in adult zebrafish, I observed animals that failed to
completely regenerate the resected bone area. There was a range of incomplete bone regeneration
adult fish that could be categorized by the degree of regenerated cartilage and bone observed in
alcian and alizarin skeletal staining. In mouse rib removal, the regenerative ability requires that
the periosteum not be removed (Srour et. al, 2016; Tripuraneni et al., 2016; Kuwahara et al.,
2019). Adult fish with no cartilage callus at either end of the resection area likely failed to
regenerate because of the inadvertent removal or damage of connective tissue specifically the
periosteum. Partially regenerated lower jawbone fish show a significant cartilage callus at both
resected ends but fail to connect and bridge the resected area. Incomplete regeneration in these
fish are likely due to the inability to stabilize the two resected bone edges with orthopedic
surgical pin implants. While it has been demonstrated that regenerative ability does decline in
zebrafish as a consequence of aging and senescence, it is unlikely that incomplete bone
regeneration is due to the zebrafish age (Tsai et al., 2007). The average age of adult zebrafish
used by Paul et al. (2016) was approximately 6 months post fertilization up to 1 year post
fertilization. In an effort to decrease the time cycle between generations, I tested the idea that the
jawbone resection surgeries could be shortened to at least 3 months post fertilization. At 7 weeks
post fertilization, and between 15-17mm standard body length, I performed lower jawbone
resection surgeries on Tubingen background and performed skeletal staining assays to observe
the patterns of bone regeneration. The younger adult fish regenerated the mandible with
improved consistency and showed a regenerative cartilage callus, cartilage bridging across the
resected area and regenerating bone all at earlier time points than demonstrated in adult fish
(Figure 3-4).
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Figure 3-4. Defining the quality of lower jaw bone regeneration in Juvenile fish
Lower jaw bone regeneration in younger fish was tested in age and size matched comparison
resection experiments. Uninjured and resected fish were measured for standard body length (SL)
less than 1 mm. Juvenile animals, 7 wpf, at 3 time points post-surgery were assessed for
regeneration qualities observed in adult animals, 6mpf. By 3 weeks post-surgery, a greater
number of juvenile animals displayed both cartilage and bone regeneration than observed in 6
mpf adults (Data not shown).
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Figure supplement 3-1. Juvenile lower jaw bone regeneration activates RUNX2+ cells for
cartilage callus formation
Juvenile fish, 7 wpf (SL 17 mm), exhibit similar bone regeneration patterns observed in adults
demonstrated in trichrome histological staining but in a shorter amount of time. Additionally,
runx2-GFP+ expressing cells are observed in the regenerative mesenchyme at 3 dpr. n = 3. By 7
dpr GFP+ cells are also observed in regenerative chondrocytes. n = 3. At 10 dpr there are GFP+
cells observed immediately adjacent to regenerated bone. n = 3. Sp7-GFP+ cells are not observed
in the regenerative mesenchyme at 3 dpr or the regenerative chondrocytes at 7 dpr. 3 dpr: n = 4.
7 dpr: n = 4. At both 7 and 10 dpr GFP+ cells are observed along the edge of regenerated bone
connected to pre-existing bone as well as regenerated bone fragments unconnected to pre-
existing bone. Dashed lines represent the 10 dpr: n = 4.
3.2.4 Lineage contribution of osteoblasts and chondrocytes to cartilage
callus regeneration
Zebrafish have an unlimited capacity to regenerate the caudal fin following amputation and that
mature osteoblasts contribute to the blastema formation of the fin rays in part through
dedifferentiation into a proliferative osteoblast progenitor state (Knopf et. al, 2011; Sousa et al.,
2011). Additionally, the unique skeletal regeneration via dedifferentiation was further
investigated by fin fracture and the dermal bone of the zebrafish skull calvarium (Geurtzen et al.,
2014). Across these skeletal sites, the adjacent mature osteoblasts underwent a dedifferentiation
to migrate to the injury site to re-differentiate back to mature osteoblasts. Given that the Crump
lab was able to obtain transgenic Cre-inducible lineage tracing lines with promoters for
osteoblasts and chondrocytes, sp7 (osterix) and col2a1. I performed lower jaw resection
surgeries on adult zebrafish to determine the contribution of pre-existing osteoblasts and
78
chondrocytes to the regenerative mesenchyme and eventually hybrid cartilage callus. In my
characterization of early time points of lower jawbone regeneration, I observed that the
transgenic labeling of osteoblasts with the sp7 promoter at 2 dpr showed a marked reduction of
GFP+ signal in the osteoblasts immediately adjacent to the resected area and a progressive
increase in GFP signal the more distal from the resected edge. If lower jawbone regenerative
response to injury was similar to what has been observed in fin rays and skull calvarium, this
result was suggestive that osteoblasts may be dedifferentiated to an earlier osteoblast progenitor
state and downregulating the expression of sp7 and migrating into the regenerative mesenchyme.
Alternatively, GFP+ cells osteoblasts could deplete signal through rapid cellular proliferation.
Four days prior to lower jaw resection surgery I treated adult sp7-CreERT2;bactin2-loxp-BFP-
stop-loxp-dsRed-stop fish with 5µM 4OH-Tamoxifen for 2 days and 2 days for drug wash out.
When I looked at 4 dpr I could observe dsRed+ derived osteoblast cells in the regenerative
mesenchyme as well as lining the surface of uninjured mineralized bone. In fact, at the proximal
end of the resected bone I observed an enrichment of dsRed+ cells and presumed de novo
regenerated bone based on the bone matrix morphology. By 8 dpr, the de novo bone connected to
pre-existing bone had increased but dsRed+ cells within the regenerative mesenchyme were not
observed. Additionally, the regenerative callus at 8 dpr were not labeled with dsRed+ cells.
Taken together, these preliminary results suggest that while some osteoblast derived cells do
move into the regenerative mesenchyme early during regeneration, there is not contribution to
the hybrid cartilage callus. It is important to note that the transgenic sp7-CreERT2 line in my
hands showed dsRed recombination in the absence of tamoxifen, however the spurious dsRed
recombination was specific to osteoblasts in the opercle and other craniofacial bones at
embryonic stages. In an abundance of caution, it is wise to consider the lower jawbone
regeneration results under the control of a constitutive Cre-promoter rather than an inducible
Cre-promoter. Any dsRed+ cells observed in the regenerative mesenchyme or on the surface of
de novo bone could have recombined to express dsRed after the injury. In the previously
described col2a1-CreERT2; bactin2-loxp-BFP-stop-loxp-dsRed-stop chondrocyte lineage trace
transgenic fish line, I treated embryos with 4OH-Tamoxifen at 5µM concentration for 12 hours
at 4 dpf to label chondrocytes with dsRed expression. As adults, I performed lower jawbone
resection surgeries at 8 wpf and an average standard body length of 17 mm. At 7 dpr, I observed
dsRed cells labeling the Meckel’s cartilage and low dsRed cells in the regenerative mesenchyme
around the surface of regenerated bone. However, due to high background the low dsRed cells
79
may represent false signal. Further experiments need to be performed to determine chondrocyte
derived contribution to the regenerative mesenchyme and hybrid cartilage callus. Thus far, I have
not observed suggestive evidence that pre-existing osteoblasts or chondrocytes are contributing
to the majority of the regenerative mesenchyme or the cartilage callus during lower jawbone
regeneration.
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Figure 3-5. Contribution of derived osteoblasts and chondrocytes to cartilage callus
regeneration
81
(A) In sp7-GFP adult fish at 2 dpr GFP+ cells were observed to progressively decrease moving
from the distal jaw joint towards the edge of the bone resected area. The contra-lateral side of the
resected lower jaw shows unaltered GFP expression. Signal intensity analysis demonstrates the
reduction of GFP expression directly adjacent to the bone but increases distally from the end of
the resection. n = 3.
(B) At 7 dpf, sp7-CreERT2;B>R embryos were treated with tamoxifen and screened for specific
dsRed expression in the opercle (OP) 18 hpt. Sibling embryos not treated with tamoxifen were
screened for errant Cre expression. n = 8. Adult lower jaw resected fish show dsRed expressing
cells lining the edge of pre-existing bone and regenerated bone is observed immediately adjacent
to pre-existing bone at 4 dpr. n = 4. Osteoblast-derived cells are observed in the regenerative
mesenchyme at 4 dpr but do not show contribution to the cartilage callus at 8 dpr. Regenerated
bone fragments at 8 dpr is observed not connected to pre-existing bone. n = 5.
(C) At 7 dpr, col2a1-CreERT2;B>R adult jaw resected fish show dsRed positive cells in the
regenerative mesenchyme. dsRed positive cells with a mesenchymal-like cell morphology are
observed in a stream of cells from the perichondrium. Perichondrium and Meckel’s cartilage are
labeled by dsRed expression prior to injury. n = 4.
3.2.5 Upper Jawbone Regeneration
Mammalian long bone fractures have been thoroughly studied, and the fracture will
endogenously develop a cartilage intermediate, called a cartilage callus, before undergoing
remodeling to replace the cartilage with bone (He et al. 2017). In short this indicates that
endochondral bones develop and heal through an endochondral or cartilage intermediate
mechanism. What is not entirely clear is if the bone formation and subsequent ossification that a
particular skeletal element experiences in a skeletal development context then inform in a
predictive way of the response to injury and regeneration. A current model of zebrafish
intramembranous skeletal regeneration indicates regeneration occurs through dedifferentiation of
osteoblasts. Specifically, this conclusion was made by Guertzen et al. via observations of
downregulation of ocn-GFP (dedifferentiation) near the injury site followed by upregulation of
runx2b and eventually upregulation again of ocn-GFP (redifferentiation). It is important to note
that these skeletal regeneration studies were performed in the caudal fin and calvaria (Guertzen
et al. 2014, Knopf et al. 2011). When I performed large-scale upper jawbone resections in the
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premaxilla of adult zebrafish, I observed a noticeably less robust regenerative cartilage callus.
Specifically, there is a small amount of alcian staining at the edges of the resected premaxilla at 7
dpr and across multiple time points out to 9 wpr, a large regenerative cartilage callus is not
observed despite the eventual complete regeneration of bone (Figure 3-6, Figure supplement 3-
2). Other than dedifferentiation of mature osteoblasts, it has been shown that the periosteum
likely houses SSCs that contribute to bone repair (He et al. 2017, Lageneste et al. 2018). To this
point, we have preliminary evidence that the cell layer immediately adjacent to the bone matrix
expresses transcripts for both col2a1 and col1a1 under a single incision of the premaxilla in adult
zebrafish (Figure supplement 3-2B). This expression pattern is reminiscent of the hybrid
cartilage callus seen in the lower jawbone regeneration. Taken together this initial result is
suggestive that a periosteal population from an intramembranous ossification origin expresses
chondrocyte-associated transcripts during a regenerative response. This indicates that
intramembranous and endochondral bones may have different processes for repair.
Figure 3-6. Pre-maxilla resection regenerates in adult zebrafish
(A) Surgical resection of the pre-maxilla on anesthetized adult zebrafish is performed by two
incisions followed by the removal of the bone fragment (1 mm) without removal of the
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connective tissue covering the bone. Alcian and Alizarin stained heads of post-surgery (0 dpr)
and uninjured fish demonstrate the pre-maxilla and not maxilla have been resected. n = 5.
(B) At 10 dpr, the pre-maxilla has regenerated bone with faint alcian stain for cartilage between
the resected bone ends. n = 6. By 9 wpr, the pre-maxilla has bridged the resected area with bone
and no cartilage is observed at the site of resection. n = 6.
(C) Dissected upper jaw including the maxilla and pre-maxilla show details of bone
regeneration. Alcian cartilage stain is observed at the edges of the resected bone at 7 dpr. By 14
dpr cartilage bridged the resected area connecting regenerated bone. Within 35 dpr no cartilage is
observed, and regenerated bone is connected across the resected area. 0 dpr: n = 4. 7 dpr: n = 6.
14 dpr; n = 6. 35 dpr: n = 6.
Figure supplement 3-2. Pre-maxilla regeneration generates cartilage while periosteal cells
co-express col2a1a and col1a1a.
(A) High magnification images of dissected pre-maxilla resected adult fish demonstrate the
patterns of cartilage and bone regeneration. In 7 dpr alcian stains cartilage extending from both
edges of the remaining bone. In 14 dpr regenerated bone fragments are connected by interspersed
regenerated cartilage. By 21 dpr cartilage is observed at the bridge of regenerated bone and at 35
dpr regenerated bone has remodeled and no cartilage is observed.
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(B) In single incision surgeries of the pre-maxilla by 10 dpr, the fluorescent in situs of
consecutive sections show co-expression of col2a1a with col1a1a along the bone surface
suggestive of periosteal cell response to bone injury. n = 3.
3.3 Discussion
As the introduction in Chapter 1 discussed the vertebrate skeleton particularly the zebrafish
craniofacial skeleton, comprises elements that not only illustrate a diversity of structure and
formation but also represent evolutionary and functional significance. In this project, myself and
Dr. Sandeep Paul worked to establish a new surgical method for large-scale bone regeneration in
the lower jawbone of adult zebrafish. We hypothesized that the unique regenerative capacity in
zebrafish caudal fin and skull calvarium would be insightful to understand in an equivalent
skeletal injury system used in mice. While the pectoral fin is the evolutionary equivalent to the
appendages in mammals, the fin rays to a degree represent digits. The mandible in zebrafish and
mammals represent a strong genetic conservation but it is important to understand that the
Meckel’s cartilage in mammals does not exist into adulthood and is thought to be absorbed and
replaced by bone. Zebrafish mandible is an advantageous skeletal element based on the ease of
accessibility, relatively large size compared to other skeletal elements, and the physical force
applied to the mandible needed for feeding can be compensated for temporarily by the jaw joint
skeletomuscular structures on the uninjured side of the mouth.
We were pleased to discover that the lower jawbone regeneration proceeds through a hybrid
cartilage bone cell population that we characterized extensively through osteo- and chondrogenic
skeletal markers. Early timepoints during regeneration suggest an enrichment of RUNX2
promoter activation in the periosteal cell layers adjacent to mineralized bone matrix and the
regenerative mesenchyme. It is unclear if a resident skeletal progenitor cell population is rapidly
expanded in response to injury or if differentiated skeletal cells activate cell fate reprogramming
to contribute to the regeneration. Either or both cell populations may contribute to the
regeneration process resulting in a heterogenous regenerative response to a large-scale skeletal
injury. Periosteal lineage tracing experiments would determine if a significant portion of the
hybrid cartilage-bone callus arises from a periosteal skeletal progenitor as has been observed in
mammalian fracture repair.
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My preliminary data performing lineage tracing experiments on pre-existing osteoblasts and
chondrocytes indicate that relatively low contribution to the regenerative mesenchyme and
hybrid cartilage callus. Osteoblasts do appear to generate new bone matrix that is attached to
uninjured bone matrix however, I observed there is a tendency for the immediate osteoblast
injury response to produce new bone matrix that covers the posterior resected area including the
remaining Meckel’s cartilage. Careful analysis will need to be performed to determine if the
uninjured Meckel’s cartilage is maintained after regeneration, but I am confident that the
Meckel’s cartilage or other chondrocyte cells are not re-established or maintained in the
regenerated area. If the Meckel’s cartilage or perichondrium are contributing a significant
portion of cells to the hybrid cartilage callus, a secondary resection would test the regenerative
capacity for repeated injury.
In an effort to reduce the skeletal cell types that could contribute to a regenerative response,
myself and Ashley Jacks developed a new surgical model in adult zebrafish by resection a
segment of the premaxilla of the upper jaw bone. The upper jaw has similar surgical advantages
as the mandible but is composed of bone and not cartilage cell types within the resected area.
The upper jaw regeneration progresses without generating a significant cartilage callus but
preliminary in situ results in a single incision injury suggest overlapping expression of col2a1
and col1a1 in the cell layers immediately adjacent to the bone matrix. Further experiments will
need to be performed on resected animals to determine if a hybrid cartilage callus is formed at
any point during regeneration. Alcian blue staining indicates some unmineralized cartilage
matrix is generated but the cellular source is not defined. To understand the contributions of
differentiated and resident progenitor skeletal cells to zebrafish skeletal regeneration, careful
lineage tracing and cell specific ablation techniques need to be employed. Cell ablation
experiments will be most effective when targeting the main source of the regenerative
mesenchyme.
3.4 Materials and Methods
3.4.1 Histology and Paraffin embedding
Skeletal staining with Alcian Blue and Alizarin Red were performed as described (Walker and
Kimmel, 2007). Following euthanasia, isolated adult heads were fixed in 4%paraformaldehyde
86
for 4 days, decalcified in 20% EDTA for 5 days and embedded in paraffin. 5μm sections were
cut on a Shandon Finesse Me+ microtome (cat no. 77500102) and collected on Apex Superior
Adhesive slides (Leica Microsystems, cat. no. 3800080). H&E staining (VWR) and trichrome
staining (Newcomer Supply) were according to the manufacturer’s instructions. Pentachrome
and Trichrome staining were performed according to manufacturer's instructions (Movat-Russell
modified pentachrome stain kit, Newcomer Supply cat. no. 9150A; Gomori One-Step, Aniline
Blue, trichrome stain kit, Newcomer Supply cat. no. 9176A).
3.4.2 Immunohistochemistry and in situ hybridization
Fluorescent in situ hybridization (ISH) on paraffin sections was carried out as described
(https://wiki.zfin.org/display/prot/3+color+Fluorescent+in+situ+on+sections). Probe templates
were amplified and cloned into pCR-BluntII-TOPO (Invitrogen), followed by linearization and
in vitro transcription following the manufacturer’s instructions (Roche Life Science,
11175025910). Immunohistochemistry was performed according to Stewart et al. (2014) with the
exception of blocking with 2% normal goat serum (Jackson Immuno Research, 005-000-121).
Primary antibodies include rabbit anti-GFP (1:500, Torrey Pines, TP401), rabbit anti-mCherry
(1:250, Novus Biologicals, cat. no. NBP2-25157), and mouse anti-GFP (1:50, Sigma Aldrich,
11814460001). Alexa Fluor secondary antibodies were used. After cutting thin sections, I
performed antigen retrieval by treating slides with citrate buffer (pH 6.0) in a steamer set (IHC
World, cat. no. IW-1102) for 35 min. I used AlexaFluor secondary antibodies and Hoechst to
visualize nuclei.
3.4.3 Imaging
Brightfield images of hematoxylin and eosin and trichrome stains were acquired with a Zeiss
AxioImager.A1 microscope and a Zeiss slide scanner AxioScan.Z1. Fluorescence images were
captured on a Zeiss LSM780 confocal microscope, with representative sections or maximum
intensity projections shown as specified. Tiling was performed using ZEN software or stitched
together using Adobe Photoshop CC. Brightness and contrast were adjusted in Adobe Photoshop
CC with similar settings for experimental and control samples.
87
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Chapter 4
Future Perspectives about Skeletal cell fate plasticity
4 Abstract
Zebrafish skeletal development and regeneration are nascent research topics that contain many
unanswered questions. While in many instances zebrafish share mechanistic and molecular
signaling patterns studied in mammalian skeletal research, there are also unique skeletal
characteristics like robust skeletal regeneration. My research has demonstrated that chondrocytes
have the plasticity to generate at least osteoblasts and adipocytes. However, the mechanisms
controlling the cell fate decisions for either lineage have not been fully defined. Skeletal cells in
vertebrates are diverse and specialized for dynamic regulation of the skeletal system however the
interplay of both immune cells and vasculature are closely involved with bone formation and
regeneration and not been fully explored. Last, the zebrafish periosteum and perichondrium
house the adult skeletal progenitor cells that contribute to bone homeostasis and regeneration. By
testing unique markers expressed early in skeletal development we may identify a strong adult
marker for further profiling to investigate questions about adult skeletal stem cell maintenance
and response to injury.
4.1 Adipogenic and osteogenic cell fate dynamics
The extent that multiple differentiated skeletal cells have the plasticity to generate different
skeletal cells raises interesting questions about the mechanisms of cell fate decisions in an injury
and non-injury context. In my work, I suggest hypertrophic chondrocytes have the ability to
participate in their release from the cartilage matrix in part by internal expression of mmp9.
These cells appear to dedifferentiate and express markers for adult skeletal stem cells. Through
processes that are not completely understood the dedifferentiated cells show contribution to
adipocyte, osteoblast, and stromal cell lineages. While the molecular signaling and cellular
interactions involved in adipogenesis are not completely understood, future investigations into
modulating molecular expression of Schnurri-2 and BMP-2 can help refine marrow adipocyte
function (Jin, 2006). Correspondingly, it would be of interest to investigate if as a consequence
92
of adipogenic signaling disruption are chondrocyte derived osteoblast contributions shifting as a
compensatory mechanism.
When I performed pentachrome on older adult zebrafish, approximately 1 ypf, I observed the
ceratohyal growth plate had reduced in area and reduced in cartilage matrix staining. I
hypothesize that if remodeling of the growth plate chondrocytes is an ongoing process
throughout aging, then potentially the cartilage matrix is progressively reduced. As a
consequence of cartilage matrix removal, the proliferative chondrocytes may differentiate into
terminal hypertrophic chondrocytes. It would be interesting to test the capacity for cell fate
plasticity to occur later in adulthood and well after bone remodeling has initiated. Performing
chondrocyte labeling experiments in zebrafish at 6 mpf or older would help define the capacity
for hypertrophic chondrocytes to contribute to bone and fat cell lineages beyond the initial
endochondral remodeling event.
In preliminary experiments, I was able to titrate tamoxifen dosing down to consistently identify
single cell labeled chondrocytes in the growth plate proliferative zone at embryonic stages. These
experiments would confirm the capacity for a single growth plate chondroblast to give rise to the
multiple skeletal cells I observe in the remodeled ceratohyal. Occasionally, I would observe
incomplete labeling within the adult growth plate, one explanation is the inefficient
recombination and variable penetrance of tamoxifen. However, the growth plate has two growth
directions and sources, interstitial and appositional, from the proliferative chondrocytes and from
the perichondrium respectively. Since the col2a1 transgenic line does not label perichondral cells
during embryonic stages it is possible the adult growth plate reflects the contribution of
proliferative chondroblasts and excludes perichondral contribution. Additionally, it would be
interesting to use the single chondrocyte labeled experiments to better understand the function of
the hypertrophic chondrocytes produced during the initial cartilage template formation,
especially at the formation of the bone collar and the start of remodeling. One could investigate
if the hypertrophic chondrocytes generated from the initial mesenchymal condensation survive
and contribute to other skeletal cell types of the remodeled bone.
Within the mmp9 mutant ceratohyal, I observe a reduction but not a complete absence of marrow
adipocytes. These marrow adipocytes may still be chondrocyte derived cells through functional
redundancy of mmp9 by other members of the mmp degradation proteins, humans have at least
93
28 Mmp related family members and zebrafish have approximately 15 mmp genes (Verma and
Hansch, 2007; Huxley-Jones et al., 2007). The extracellular matrix does not serve solely as a
structural scaffolding function but also functions to provide cell signaling and has been shown to
have roles in differentiation decisions (Ishizeki, 2012). Therefore, the expansive family of mmp
proteinases with specialized function and expression patterns may hold insights into mechanisms
of skeletal cell plasticity.
4.2 Invading immune and vasculature cells dynamics with
skeletal cells
The bone remodeling event in the endochondral ceratohyal is not an external injury but may use
systemic cells like immune and vasculature to stimulate remodeling as observed during fracture
repair. Matrix absorbing cells like osteoclasts and chondroclasts were not investigated for bone
regeneration or development. In addition to the expression of cartilage ECM degradation
proteins, the involvement of chondroclasts as a cellular source for hypertrophic chondrocyte
release is a point of interest. In mice, the endochondral bone surface is thought to be lines with
multi-nucleated osteoclasts while osteoblasts line the periosteal surface (Hall, 2015; Knowles et
al., 2012). In my study of how the ceratohyal bone undergoes endochondral remodeling,
zebrafish skeletal bone maintenance research was lacking. While zebrafish have osteoblasts,
osteoclasts, chondroblasts and chondroclasts, there is not a clear understanding if zebrafish bones
perform homeostasis in what has been observed in mammalian systems.
While I characterized the increasing vascularization of the ceratohyal, I did not test the
requirement of vascular cells for the hypertrophic chondrocyte remodeling event. Mouse
endochondral bone remodeling requires the involvement and signaling of vasculature endothelial
cells (Ramasamy et al., 2014). It is unclear if systemic cells or vasculature cells interact with the
release of hypertrophic chondrocytes or their subsequent cell fate decisions. In an effort to
understand how the inflammatory response early in regeneration can induce the skeletal
progenitor cells or the cell fate plasticity of differentiated skeletal cells, I have gathered
transgenic zebrafish labeling cells of the immune system as well as mutant lines with
demonstrated defects in early immune development. These transgenic lines can be used trace
94
macrophages and neutrophils as well as conditionally ablated through a metronidazole-
nitroreductase system. Recently, a zebrafish SCID transgenic line was established through
mutations in the Prkdc gene has been characterized to not have functional T and B lymphocytes
(Jung et al., 2016). To date the effects of immunodeficiency on skeletal development and
endochondral bone remodeling have not been described.
4.3 Opportunities to define perichondral and periosteal cell
populations
The perichondrium and eventual periosteum are established during and shortly after
mesenchymal condensation. The perichondrium is not formed directly through chondrogenesis
(Colnot et al., 2004). There are cell to cell interaction proteins that are unique to the
perichondrium during mesenchymal condensation and skeletal chondrogenesis like Tenascin C,
Syndecan, and NCAM. Syndecans are a family of transmembrane proteins that in conjunction
with heparin sulfate and chondroitin sulfate chains build glycoaminoglycans and are involved in
growth factor activation, matrix adhesion, and cell to cell adhesions (Lambaerts et al., 2009).
Syndecans in the extracellular matrix interact with collagens, thrombospondin and tenascin
(Morgan et al., 2007). There are 3 Syndecan genes in zebrafish and in mouse embryonic studies
Syndecan 2 and 3 expression has been observed in the presumptive perichondral layer during
mesenchymal condensations of skeletal elements (Koyama et al., 1996; David et al., 1993).
Adult zebrafish Syndecan expression has not been explored as a potential marker of periosteum
and perichondrium. Tenascin C and tenascin W is expressed in mouse embryonic periosteum of
the rib (Scherberich et al., 2003; Chiquet-Ehrismann and Tucker, 2004). In in vitro conditions,
osteoblast-like cells respond to mechanical loading with increased tenascin C expression in rat
ulnae (Webb et al., 1997). Importantly, Geurtzen et al. (2014) observed fibroblasts and
osteoblasts expressing Tenascin C in zebrafish ray fractures 1 day post injury. It could be
interpreted the expanded expression of Tenascin C in the fibroblasts adjacent to the osteoblasts
are skeletal progenitor cells within the periosteum responding to the skeletal injury.
95
4.4 References
Chiquet-Ehrismann, R., and Tucker, R.P. (2004). Connective tissues: signalling by tenascins. Int.
J. Biochem. Cell Biol. 36, 1085–1089.
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(2014). Mature osteoblasts dedifferentiate in response to traumatic bone injury in the zebrafish
fin and skull. Development 141, 2225–2234.
Hall, B.K. (2015). Chapter 24 - Osteoblast and Osteocyte Diversity and Osteogenesis In Vitro. In
Bones and Cartilage (Second Edition), B.K. Hall, ed. (San Diego: Academic Press), pp. 401–
413.
Huxley-Jones, J., Clarke, T.-K., Beck, C., Toubaris, G., Robertson, D.L., and Boot-Handford,
R.P. (2007). The evolution of the vertebrate metzincins; insights from Ciona intestinalis and
Danio rerio. BMC Evolutionary Biology 7, 63.
Ishizeki, K. (2012). Imaging analysis of osteogenic transformation of Meckel’s chondrocytes
from green fluorescent protein-transgenic mice during intrasplenic transplantation. Acta
Histochemica 114, 608–619.
Jin, W., Takagi, T., Kanesashi, S., Kurahashi, T., Nomura, T., Harada, J., and Ishii, S. (2006).
Schnurri-2 Controls BMP-Dependent Adipogenesis via Interaction with Smad Proteins.
Developmental Cell 10, 461–471.
Jung, I.H., Chung, Y.-Y., Jung, D.E., Kim, Y.J., Kim, D.H., Kim, K.-S., and Park, S.W. (2016).
Impaired Lymphocytes Development and Xenotransplantation of Gastrointestinal Tumor Cells in
Prkdc-Null SCID Zebrafish Model. Neoplasia 18, 468–479.
Knowles, H.J., Moskovsky, L., Thompson, M.S., Grunhen, J., Cheng, X., Kashima, T.G., and
Athanasou, N.A. (2012). Chondroclasts are mature osteoclasts which are capable of cartilage
matrix resorption. Virchows Arch 461, 205–210.
96
Koyama, E., Shimazu, A., Leatherman, J.L., Golden, E.B., Nah, H.D., and Pacifici, M. (1996).
Expression of syndecan-3 and tenascin-C: possible involvement in periosteum development. J.
Orthop. Res. 14, 403–412.
Lambaerts, K., Wilcox-Adelman, S.A., and Zimmermann, P. (2009). The signalling mechanisms
of syndecan heparan sulphate proteoglycans. Curr Opin Cell Biol 21, 662–669.
Morgan, M.R., Humphries, M.J., and Bass, M.D. (2007). Synergistic control of cell adhesion by
integrins and syndecans. Nat. Rev. Mol. Cell Biol. 8, 957–969.
Ramasamy, S.K., Kusumbe, A.P., Wang, L., and Adams, R.H. (2014). Endothelial Notch activity
promotes angiogenesis and osteogenesis in bone. Nature 507, 376–380.
Scherberich, A., Tucker, R.P., Samandari, E., Brown-Luedi, M., Martin, D., and Chiquet-
Ehrismann, R. (2004). Murine tenascin-W: a novel mammalian tenascin expressed in kidney and
at sites of bone and smooth muscle development. Journal of Cell Science 117, 571–581.
Verma, R.P., and Hansch, C. (2007). Matrix metalloproteinases (MMPs): Chemical–biological
functions and (Q)SARs. Bioorganic & Medicinal Chemistry 15, 2223–2268.
Webb, C.M.B., Zaman, G., Mosley, J.R., Tucker, R.P., Lanyon, L.E., and Mackie, E.J. (1997).
Expression of Tenascin-C in Bones Responding to Mechanical Load. Journal of Bone and
Mineral Research 12, 52–58.
Abstract (if available)
Abstract
The skeletal system as an organ develops and regenerates through complex interactions with surrounding tissues and vasculature. Cellular components of bone, cartilage, and fat all demonstrate cell fate plasticity to contribute to the other skeletal cell lineages. Importantly, the physical surroundings especially the building, breakdown, and remodeling of the matrix components of bone, cartilage and fat are involved in the differentiation and dedifferentiation of skeletal cells. While the periosteum is vital to fracture repair healing, the origins of and unique molecular markers of the skeletal progenitor cell population are not yet understood. Using zebrafish as a model for both skeletal development and regeneration can unravel the complex interactions of many skeletal cell types while defining the induced or inherent skeletal cell fate plasticity seen across vertebrates. ❧ Much of the vertebrate skeleton develops from cartilage templates that are progressively remodeled into bone. Lineage tracing studies in mouse suggest that chondrocytes within these templates persist and become osteoblasts, yet the underlying mechanisms of this process and whether chondrocytes can generate other derivatives remain unclear. I find that zebrafish cartilages undergo extensive remodeling and vascularization during juvenile stages to generate fat-filled bones. Growth plate chondrocytes marked by sox10 and col2a1a contribute to osteoblasts, marrow adipocytes, and mesenchymal cells within adult bones. At the edge of the hypertrophic zone, chondrocytes re-enter the cell cycle and express leptin receptor (lepr), suggesting conversion into progenitors. Further, mutation of matrix metalloproteinase 9 (mmp9) results in delayed growth plate remodeling and fewer marrow adipocytes. My data support Mmp9-dependent growth plate remodeling and conversion of chondrocytes into osteoblasts and marrow adipocytes as conserved features of bony vertebrates.
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Asset Metadata
Creator
Giovannone, Dion Francis
(author)
Core Title
Skeletal cell fate plasticity in zebrafish bone development and regeneration
School
Keck School of Medicine
Degree
Doctor of Philosophy
Degree Program
Development, Stem Cells and Regenerative Medicine
Publication Date
07/30/2020
Defense Date
04/24/2019
Publisher
University of Southern California
(original),
University of Southern California. Libraries
(digital)
Tag
adipocyte,bone,cartilage,chondrocyte,Development,jawbone,OAI-PMH Harvest,osteoblast,perichondrium,periosteum,Regeneration,Skeleton,zebrafish
Format
application/pdf
(imt)
Language
English
Contributor
Electronically uploaded by the author
(provenance)
Advisor
Mariani, Francesca (
committee chair
), Crump, Gage (
committee member
), Merrill, Amy (
committee member
)
Creator Email
dgiovann@usc.edu,dion.giovannone@gmail.com
Permanent Link (DOI)
https://doi.org/10.25549/usctheses-c89-202064
Unique identifier
UC11663288
Identifier
etd-Giovannone-7706.pdf (filename),usctheses-c89-202064 (legacy record id)
Legacy Identifier
etd-Giovannone-7706.pdf
Dmrecord
202064
Document Type
Dissertation
Format
application/pdf (imt)
Rights
Giovannone, Dion Francis
Type
texts
Source
University of Southern California
(contributing entity),
University of Southern California Dissertations and Theses
(collection)
Access Conditions
The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the a...
Repository Name
University of Southern California Digital Library
Repository Location
USC Digital Library, University of Southern California, University Park Campus MC 2810, 3434 South Grand Avenue, 2nd Floor, Los Angeles, California 90089-2810, USA
Tags
adipocyte
cartilage
chondrocyte
jawbone
osteoblast
perichondrium
periosteum
zebrafish