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Development and evolution of skeletal joints: lessons learned from studying zebrafish
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Development and evolution of skeletal joints: lessons learned from studying zebrafish
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DEVELOPMENT AND EVOLUTION OF SKELETAL JOINTS:
LESSONS LEARNED FROM STUDYING ZEBRAFISH
by
Amjad Askary
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(Genetics, Molecular, and Cellular Biology)
December 2016
© Copyright 2016 Amjad Askary
ii
This thesis is dedicated to my wife, Niki
for her support, compassion, and patience,
and to zebrafish for being an endless
source of amazement and wonder for me.
iii
Acknowledgments
I would like to first thank my mentor Gage Crump for giving me the opportunity to work in
his lab, providing a wonderful research environment, and supporting me all through the graduate
school. Thanks for letting me try out my ideas, and for showing me the path to good research.
I would also like to thank my Dissertation committee members: Neil Segil, Andrew
McMahon, and Cheng-Ming Chuong for all their help and advice throughout my career at USC.
I am also very grateful to my first mentor at USC, Michael Lieber, for his unbelievable help
and support. I cannot thank him enough for all the opportunities he provided me with, all his
thoughtful advices, and all the good times I spent in his lab.
In addition, I would like to thank everyone in Crump lab, past and present. They made my
PhD a pleasant and memorable experience. Especially, Megan Matsutani for teaching me how to
work with zebrafish embryos and helping me whenever I needed it. Simone Schindler for
patiently answering all my questions about in situ hybridization. Paul Bump for going above and
beyond in helping me with my experiments and lab duties. Joanna Smeeton, who shares my
fascination with joints, for being an amazing colleague and making our shared project a
successful endeavor. And the rest of my labmates: Samuel Cox, Bartosz Balczerski, Elizabeth
Zuniga, Ankita Das, Sandeep Paul, Lindsey Barske, Pengfei Xu, Camilla Teng, and Dion
Giovannone for all the fun times in the lab.
In addition to my labmates, I had the opportunity to collaborate, learn from, and work
alongside researchers from other labs in our department. From those, I would like to especially
thank Xinjun He, from McMahon lab, for taking so much of his time teaching me how to do gel
shift assays, troubleshooting my experiments, and helping me with the revision of my paper. I
also want to thank Lick Lai for helping me with the mouse work and for our helpful discussions,
as well as Audrey Izuhara and Suhasni Gopalakrishnan for their assistance with our in vitro
experiments.
iv
One of my most remarkable experiences during these years has been attending the
Embryology course at Marine Biological Laboratories in Woods Hole, MA. So special thanks to
all those who helped make it happen. Starting from Gage who supported me in taking the course,
the course directors Richard Behringer and Alejandra Sanchez Alvarado, all our teachers,
lecturers, TAs, and CAs in the course, and all of my classmates in Embryology 2015.
I would also like to thank all my friends in the US and all my family in Iran for always being
there for me. Thankfully, I have so many that I can’t list them all here individually by name.
Last but certainly not least, I would like to thank my wonderful wife, Niki Bayat, for her
love and support in all these years.
v
Table of Contents
Acknowledgments.......................................................................................................................... iii
List of Figures and Tables.............................................................................................................. vi
Summary ...................................................................................................................................... viii
Chapter 1 Introduction .....................................................................................................................1
Chapter 2 Iroquois Proteins Promote Skeletal Joint Formation by Maintaining Chondrocytes
in an Immature State .................................................................................................................33
Chapter 3 Unbiased search for novel regulators of joint specification in zebrafish facial
skeleton .....................................................................................................................................63
Chapter 4 Ancient origin of lubricated joints in bony vertebrates .................................................80
Conclusion ...................................................................................................................................108
vi
List of Figures and Tables
Chapter 1:
Figure 1. 1. Histological features of joints. ..................................................................................... 3
Figure 1. 2. Skeletal joints in zebrafish and mouse. ....................................................................... 7
Figure 1. 3. Distinct modes of interzone formation. ....................................................................... 9
Figure 1. 4. Iterative segmentation of the bony rays of the zebrafish fin. .................................... 12
Figure 1. 5. Critical thresholds of Bmp and Tgfβ signaling in joint development. ...................... 17
Figure 1. 6. Chondrocyte fate decisions........................................................................................ 18
Figure 1. 7. Synovial specializations. ........................................................................................... 22
Chapter 2:
Figure 2. 1. Regulation of irx7 and irx5a expression and character of the hyoid joint. ................ 36
Figure 2. 2. Requirements of Irx7 and Irx5a in hyoid joint formation. ........................................ 38
Figure 2. 3. Irx7 represses chondrocyte maturation. ..................................................................... 39
Figure 2. 4. Irx genes directly repress a col2a1a enhancer. .......................................................... 40
Figure 2. 5. Progressive restriction of irx7:GFP to the hyoid joint and Hand2-independent
inhibition of irx7 expression by Bmp signaling. ........................................................................... 44
Figure 2. 6. Hyoid joint and symplectic cartilage defects in independent null alleles for irx7 and
irx5a. ............................................................................................................................................. 47
Figure 2. 7. Potential evolutionary history of vertebrate Iroquois genes. ..................................... 49
vii
Figure 2. 8. Potential roles of Irx genes at multiple joints. ........................................................... 50
Chapter 3:
Figure 3. 1. Bmp, Edn1, and Jagged-Notch signaling pattern the facial skeleton of zebrafish. ... 66
Figure 3. 2. Transgenic lines labeling different regions of the pharyngeal arch help isolate
specific populations. ..................................................................................................................... 70
Figure 3. 3. Intermediate specific expression of the candidate genes can be verified by RNA in
situ hybridization. ......................................................................................................................... 75
Table 3. 1. List of genes enriched in the intermediate domain of zebrafish pharyngeal arches. .. 73
Chapter 4:
Figure 4. 1. Synovial-like morphology of jaw and fin joints in ray-finned fish. .......................... 84
Figure 4. 2. Live imaging of jaw joint cavitation. ........................................................................ 85
Figure 4. 3. Expression of Prg4 genes in articular chondrocytes of ray-finned fish..................... 89
Figure 4. 4. Gene expression within the zebrafish and stickleback jaw joints. ............................ 90
Figure 4. 5. Evolution of vertebrate Proteoglycan 4 (Lubricin). .................................................. 91
Figure 4. 6. Progressive deterioration of the jaw joint in zebrafish lacking prg4b. ...................... 94
Figure 4. 7. Serial sections through a representative wild-type and prg4a-/-; prg4b-/- mutant jaw
joint. .............................................................................................................................................. 96
Figure 4. 8. OARSI scoring system for zebrafish. ........................................................................ 97
Figure 4. 9. Requirement of prg4 gene function for fin joints of zebrafish. ................................. 98
viii
Sum mar y
We owe the flexibility of our bodies to the joints that connect our bones to each other.
Proper development of these joints requires the integration of multiple tissue types, made
possible by coordination of several signaling and regulatory pathways. Conceptually, the process
of joint development can be divided into three steps: positioning of the joint within the skeleton,
specification of joint cell identities, and formation of joint specializations. Positioning the
prospective joint region involves establishment of an “interzone” region of joint progenitor cells
within a nascent cartilage condensation. Inside the interzone, tight regulation of several signaling
pathways, most notably Bmp and Tgf , guides the joint chondrocytes through their cell fate
decisions and prevents their hypertrophic maturation. Joint cells then acquire further
specializations according to the type and function of each joint.
The majority of research on joint development, so far, has been conducted using mice and
chicks as model organisms. We believe that zebrafish can offer a complementary reductionist
approach towards understanding joint development. As an example, we used the hyoid joint of
zebrafish, which functions in gill ventilation, to gain new insights into the regulation of cell
identity in joints. At the hyoid joint, prospective joint chondrocytes express SoxE transcription
factors, sox9 and sox10. However, in contrast to the differentiating chondrocytes in cartilage
condensations, joint chondrocytes make only low levels of Col2a1a and Aggrecan. We found
that members of Iroquois transcription factors, irx7 and irx5a, are expressed in the hyoid joint
chondrocytes. We then generated irx7 and irx5a null mutants and showed that mutant hyoid joint
chondrocytes inappropriately mature into high Col2a1a-expressing chondrocytes, in the absence
of direct repression of a col2a1a enhancer by Iroquois proteins. In mammals, Irx1 and Irx2 are
expressed in the developing interphalangeal joints. We showed that either mouse Irx1 or
ix
zebrafish Irx7 can repress the chondrogenic differentiation of a murine chondrogenic cell line.
This finding suggests that the function of Irx genes in inhibiting excessive differentiation of joint
cells may be conserved from zebrafish to mouse.
Additionally, we took an unbiased genomic approach to find more regulators of cell fate
specification and patterning within zebrafish facial skeleton. The Facial skeleton is formed by
neural crest derived cells of the pharyngeal arches. Prior to condensation of arch mesenchymal
cells, they are patterned into at least three domains along the Dorsal-Ventral (DV) axis. Whereas
dorsal and ventral domains of the arches give rise to dorsal and ventral elements of the facial
skeleton, joints are formed by cells from the intermediate domain. We used transgenic zebrafish
lines that label different domains, or combinations of domains, within pharyngeal arches to sort
and sequence cells, to identify their transcriptome. We then used our expression data to search
for genes that are enriched in each domain. Our assumption is that the genes which are important
for specification of each domain must be expressed preferentially in those domains. This analysis
has provided us with novel candidates, for in depth genetic and functional analysis.
At birth, major joint defects are relatively uncommon. However, the progressive
degeneration of joints in osteoarthritis is the leading cause of disability in the United States.
Consequently, among all the different types of joints, the study of synovial joints, the type that is
prone to osteoarthritis, is arguably of highest clinical relevance. Synovial joints, such as those in
our limbs, are the most sophisticated and most flexible types of joints. They are composed of
multiple tissue types: permanent cartilage that cushions the articulating bones, synovial
membranes that enclose a lubricating fluid-filled cavity, and a fibrous capsule and ligaments that
provide structural support. It is commonly believed that synovial joints evolved when vertebrates
transitioned from water to land, because lubrication could facilitate movement of their load
x
bearing joints which were exposed to increased gravitational pressure on land. Consistently, at
the stage that they are best studied, i.e. early larval stages, none of the zebrafish joints resemble
the lubricated synovial joints of mammals closely. However, we provide substantial evidence
that certain joints in zebrafish body, including the jaw joint, start to acquire features of synovial
joints at later stages. Building on histological observations that the jaw joints of diverse fish
species (e.g. spotted gar, stickleback, and zebrafish) have cavities, synovial membranes, articular
cartilage, and fibrous capsules, we showed that these joints also produce Prg4/Lubricin, a
lubricating protein which also serves as a marker of synovial joints. Further, we generated
mutant zebrafish lacking the Lubricin-encoding gene, prg4b, and showed that they develop
arthritic changes to their jaw joint that closely resemble those observed in mutant mice and
human patients lacking Lubricin. Synovial properties in zebrafish are not limited to the jaw joint,
as joints at the base of the pectoral fin also produce and require Lubricin for their maintenance.
These findings show that synovial joints were present in the common ancestor of all bony
vertebrates. Particularly, the Lubricin mutant establishes the first genetic arthritis model in
zebrafish, enabling the study of synovial joints in this powerful and highly regenerative model.
Future work in zebrafish has the potential to address unanswered questions such as the cellular
mechanism of cavitation and the ability of synovial joints to regenerate.
1
Chapter 1
Introduction
2
Joints connect neighboring bones within the vertebrate skeleton. Based on their structure,
joints can be classified into three types (Figure 1.1). Fibrous joints called sutures separate the flat
bones of the skull and allow for flexibility during childbirth. Cartilaginous joints, for example
between the vertebrae, provide limited mobility and act as shock absorbers. The most
sophisticated types of joints are the lubricated synovial articulations, such as those of the highly
mobile limbs and jaw. Despite their differing structures and extents of mobility, all three types of
joints share key features. For example, developing joints are enriched for skeletal progenitor
cells, which in the case of sutures provide a continuing supply of osteoblasts for bone growth
(Gunnell et al., 2010; Kahn et al., 2009; Kozhemyakina et al., 2015; Ray et al., 2015; Zhao et al.,
2015). Conversely, each joint is characterized by its own specialized morphology and function,
which is reflected by unique patterns of gene expression by their resident skeletal and supporting
cells.
3
Figure 1. 1. Histological features of joints.
(A) Sutures are a type of immoveable articulation between bones, with the suture mesenchyme housing
progenitors for bone growth and repair. Hematoxylin and eosin staining shows comparable coronal
sutures of an E16.5 mouse embryo and young adult zebrafish. (B) The intervertebral discs of mouse and
zebrafish have a very different structure. In mammals (shown here for postnatal day 15 mouse), a
cartilage endplate covers each vertebrae, with the disc consisting of a ring of annulus fibrosus (AF) tissue
and a core of nucleus pulposus (NP) tissue. In adult zebrafish, vertebral bones (red) are separated by
layers of fat that appear white upon sectioning. (C) Synovial joints are freely moveable articulations
characterized by fluid-filled cavities lined by articular hyaline cartilage. Some synovial joints, as shown in
a section of the knee joint from an adult mouse, include additional specializations such as menisci. The
adult zebrafish jaw joint has a clear articular cartilage layer and synovium but no mensicus. In chapter 4,
we further discuss synovial characteristics of the adult zebrafish jaw joint. Photos courtesy of Camilla
Teng (A), Jennifer Zieba (B, mouse), and Denis Evseenko (C, mouse).
4
A critical early event in the creation of all joints is the suppression of osteoblast or
chondrocyte differentiation at particular sites within the developing skeleton. For many synovial
joints, suppression of chondrocyte maturation occurs within a broad region called the interzone
(Decker et al., 2014; Longobardi et al., 2015). Lineage analysis demonstrates that interzone cells
contribute not only to the articular cartilage lining the joint cavity but also to the synovial
membrane, menisci, and ligaments (Koyama et al., 2008; Rountree et al., 2004). While much
remains to be learned about how joints are positioned, the precise spatial deployment of
activators and inhibitors of diverse signaling pathways is a recurrent theme. One consequence of
such signaling is the induction of several types of transcription factors and chromatin modifiers,
which function to inhibit cartilage maturation within the interzone. What is less clear is how later
events in joint development are regulated, such as production of the lubricating protein
Prg4/Lubricin by articular chondrocytes (Ikegawa et al., 2000; Rhee et al., 2005), creation of the
joint cavity (Archer et al., 2003), and the local differentiation of interzone cells into menisci and
ligament attachment points (entheses).
While major congenital anomalies of joints are uncommon, the progressive loss of joint
structure and function in osteoarthritis is the leading cause of disability in the United States
(Helmick et al., 2008). Genetic factors contribute substantially to osteoarthritis (e.g. estimated at
60% for the hip (MacGregor et al., 2000)), with recent studies beginning to uncover specific loci
linked to arthritis predisposition (Consortium et al., 2012). An emerging theme is that minor
defects in developmental morphogenesis may lead to architectural defects in postnatal joints
(Baker-LePain and Lane, 2010), which result in the increased wear-and-tear that predisposes to
osteoarthritis. A key feature of osteoarthritis is the ectopic hypertrophic differentiation of
articular chondrocytes at the joint surface, which erodes the cartilage cushions protecting the
bones and eventually the bones themselves (Pitsillides and Beier, 2011). This pathological
5
observation suggests that a challenge for prolonged joint health is preventing joint-lining
chondrocytes from undergoing hypertrophic maturation. Interestingly, many of the same
signaling pathways and transcription factors that help to establish articular chondrocyte fate
continue to be expressed at joints into adulthood, with postnatal genetic manipulations in some
cases establishing continued requirements in maintaining articular fate (Chen et al., 2008;
Rountree et al., 2004). There are also potential roles for epigenetic modifications in establishing
long-term repression of the hypertrophic program in articular chondrocytes (Zhang and Wang,
2015), as well as other factors selectively required for the maintenance but not development of
joints.
In this chapter, we first introduce zebrafish as an emerging model for the study of joint
development. We then review the existing literature on the development of skeletal joints, a
process that we conceptually break into three major steps: positioning the joint domain,
controlling chondrocyte identity at the joint, and generation of joint specializations.
1.1 Use of Zebrafish for Developmental Studies of Joints
Although most studies on joints have been conducted in mammals, the zebrafish has
recently emerged as an alternative model to investigate basic joint biology. Zebrafish have many
of the same types of joints found in mammals (Figure 1.1). Sutures connect the skull bones in
both fish and mammals, with undifferentiated mesenchyme residing between the interleaved
bony plates (Quarto and Longaker, 2005) (Figure 1.1A and Figure 1.2C,D). Imaging studies in
living zebrafish have revealed the stepwise formation of wild-type sutures, as well as abnormal
suture development upon genetic or pharmacological perturbations (e.g. loss of retinoic acid
6
signaling) (Jeradi and Hammerschmidt, 2016; Kague et al., 2016; Laue et al., 2011). Zebrafish
also have intervertebral discs that are prone to degeneration upon aging (Hayes et al., 2013).
However, the discs of adult zebrafish differ in structure from those in mammals in lacking
cartilage, with histology showing adipocytes between the vertebral bones rather than nucleus
pulposus and annulus fibrosus structures as in mammals (Figure 1.1B and Figure 1.2C,D). In the
zebrafish fry, there are two major cartilaginous joints in the head: the jaw joint between the
Meckel’s and palatoquadrate cartilages and the bipartite hyoid joint between the ceratohyal,
interhyal, and hyosymplectic cartilages (Figure 1.2A). Studies of these joints have revealed
conserved roles for GDF family ligands (Reed and Mortlock, 2010) as well as new roles for
Nkx3.2 (Miller et al., 2003) transcription factor in joint development.
7
Figure 1. 2. Skeletal joints in zebrafish and mouse.
(A) In zebrafish fry at 5 days post-fertilization, well-studied cartilaginous joints (arrows) include the jaw
joint between Meckel’s and palatoquadrate cartilages and the bipartite hyoid joint between
hyosymplectic, interhyal and ceratohyal cartilages. (B) In a mouse embryo at E17.5, representative joints
include the shoulder and elbow joints (arrows) and interphalangeal joints (arrowheads). As in young
zebrafish, these joints are largely cartilaginous at this stage. (C) The adult zebrafish skeleton is largely
composed of bone and contains many types of joints (arrows), including sutures in the skull,
intervertebral joints, and synovial-like joints in the jaw and pectoral fin. (D) The mouse skeleton at
postnatal day 21 has similar joints to zebrafish, including sutures, intervertebral discs, and synovial joints
8
in the knee and digits (i.e. interphalangeal). ch, ceratohyal; dr, distal radial; fe, femur; hs, hyosymplectic;
ih, interhyal; M, Meckel’s; pq, palatoquadrate; pr, proximal radial; t, tibia.
1.2 Positioning the Joint Domain
The future joint domain could be determined in at least two major ways (Figure 1.3).
First, an initial chondrogenic condensation can be split into two or more distinct zones by
creation of a morphologically distinct interzone region, which is characterized by flattened and
highly compacted cells relative to the adjacent growth plates. An example of this mechanism is
the creation of the interphalangeal joints in the digits. Second, the prospective joint domain can
be created by the appositional growth of two neighbouring condensations, such as what occurs
between the anlagen of the femur and pelvic bone (Capellini et al., 2011) and for the temporal-
mandibular joint of the mammalian jaw (Purcell et al., 2009). Interestingly, compromised
condensation formation in barx1 zebrafish mutants can lead to ectopic joints, potentially through
aberrant condensation splitting (Nichols et al., 2013). It remains unclear, however, what effects
the mode of joint specification has on later development. In principle, the preservation of
progenitor zones at the leading edges of appositionally growing condensations may result in their
later fusion into a structure closely resembling an interzone.
9
Figure 1. 3. Distinct modes of interzone formation.
The joint interzone develops in the position of the presumptive joint and precedes articular cartilage
differentiation and joint cavitation. The interzone can be generated from a single mesenchymal
condensation (A, green) or through appositional growth of adjacent condensations (B). While cells
outside the interzone undergo further cartilage differentiation (blue) and eventually hypertrophy and
mineralize during endochondral bone development (red), cells within the interzone are maintained as
immature chondrocytes at the articular surface (green flattened cells) and contribute to joint
specializations such as the synovial membrane (orange). Also shown are chondrocyte progenitors within
10
the perichondrium (blue flattened cells) and osteoblast progenitors within the periosteum (red flattened
cells).
In many cases, joints develop in a periodic pattern, e.g. the interphalangeal joints of the
digits. Studies of the ray joints of the zebrafish fins have begun to uncover potential mechanisms
of iterative joint specification. The proximal portion of the fish fin is thought to be homologous
to tetrapod limbs (Tanaka, 2016), with the more derived distal portion containing a series of
bony rays segmented at regular intervals by fibrous joints (Figure 1.4). While these ray joints
differ from the synovial-like joints at the base of the fin, they have been useful for understanding
how repeated joints can be established. Loss-of-function mutations in the gap junction protein-
encoding gene connexin43 (short-fin) result in joints spaced closer together, independent of
effects on fin size, with Connexin43 overexpression eliminating joints (Hoptak-Solga et al.,
2007; Sims et al., 2009). Gain-of-function mutations in the potassium channel-encoding gene
kcnk5b (another-long-fin) also result in fins with irregularly spaced joints (Perathoner et al.,
2014). In addition, the even-skipped transcription factor evx1 is essential for ray joint
development in zebrafish (Borday et al., 2001; Schulte et al., 2011; Ton and Iovine, 2013). While
it is unclear the extent to which iterative patterning of ray joints would be relevant to mammalian
joint patterning, mesenchymal cells within the interzone of developing mammalian joints do
express Connexins 32, 40, and 43 (Archer et al., 2003; Schlegel et al., 2009), and Connexin40
-/-
mice display losses of specific joints in the distal limbs (Pizard et al., 2005). Mutations of
CONNEXIN43 in Oculodentodigital Dysplasia syndrome also result in skeletal defects, including
missing and fused phalanges (Paznekas et al., 2009). The conserved requirement of Connexins in
fish and mammals suggest that gap junctions could be required for the efficient spread of some
unknown small molecule between cells in both species, although most growth factors would be
11
assumed to be too large to pass through these channels. Nonetheless, modelling studies in
zebrafish reveal how the expression and diffusion of just two factors is sufficient to explain the
repeated spacing of joints within the fin (Rolland-Lagan et al., 2012). At the tip of the elongating
fin, a growth factor would promote mesenchymal proliferation and inhibit joints, with joint
formation being induced when the concentration of the growth factor falls below a certain
threshold. At this position, a joint-promoting factor would be induced, which would secondarily
inhibit other joints from forming nearby. Only in positions where the two factors were below a
critical threshold would a new joint form (Figure 1.4B). It is tempting to speculate that a similar
mechanism might occur for joints in the hands and feet of tetrapods, such as the periodically
spaced phalangeal joints of the digits. Good candidates for the limb/fin growth factor are Fgfs,
which are produced by the apical ectoderm ridge and required for limb/fin outgrowth (Martin,
1998). Wnt9a/14 is a candidate joint factor that would inhibit additional joints at a distance, as its
misexpression in mouse induces an ectopic joint while inhibiting development of the adjacent
normal joint (Hartmann and Tabin, 2001). Variations on the theme of diffusible activators and
inhibitors generating repeated patterns are common in biological systems. Such reaction-
diffusion mechanisms proposed by Turing (Turing, 1952) have been used to explain the
patterning of teeth (Salazar-Ciudad and Jernvall, 2010), ectodermal appendages (Chuong et al.,
2013), and skin pigmentation (Kondo and Miura, 2010). Future work is needed to explore the
relative contribution of pre-patterning versus iterative activator-inhibitor interactions for the
positioning of different joints.
12
Figure 1. 4. Iterative segmentation of the bony rays of the zebrafish fin.
(A) The tail fin from a one-month old zebrafish was stained with alizarin red and alcian blue to highlight
bone and cartilage, respectively. In the bony rays, a series of joints form in a segmental pattern from the
base of the fin to the tip (left to right in this image). (B) Modelling predicts that just two morphogens can
generate the segmental pattern of joints in the fin rays. Bone cells are shown in red, joint cells in black
with blue nuclei, and progenitors in grey. At forming joints, one morphogen (green) specifies joint fates
while inhibiting neighboring cells from becoming joints. At the distal end, a growth factor (red) drives
progenitor growth (right arrow) that lengthens the fin while inhibiting joint formation. Over time (top to
bottom), new joints form where both morphogens fall below a critical threshold. (C) The gap junction
protein Connexin43, the potassium channel Kcnk5b, and the transcription factor Evx1 have all been
shown to be required for correct joint spacing in the fin rays. One possibility is that secreted morphogens,
as well as the transport of small molecules and electrical signaling between neighboring cells, combine to
regulate joint spacing.
13
In contrast to the repeated joints in appendages, there are other contexts where the joint
interzone appears to be pre-specified by the convergence of multiple developmental signalling
pathways. An illustrative example is the zebrafish jaw joint, which is positioned by integration of
Edn1 and Bmp signalling within the neural crest-derived precursors of the jaw skeleton
(Medeiros and Crump, 2012). In zebrafish, loss of the Edn1 ligand or its Ednra receptors results
in defects in the lower jaw and jaw joint (Miller et al., 2000; Nair et al., 2007). The joint seems
to be particularly dependent on optimal Edn1 signaling as partial reductions of the pathway in
furina, plcb3, or mef2ca mutants have greater effects on joint versus lower jaw development
(Miller et al., 2007; Walker et al., 2006; Walker et al., 2007). An important target of Edn1
signalling in joint formation is the transcription factor nkx3.2/bapx1, as antisense reduction in
zebrafish results in specific loss of the jaw joint (Miller et al., 2003). In chick, ectopic expression
of Bmp4 and Fgf8 abrogates Nkx3.2 expression and results in loss of joint structures such as the
retroarticular process (Duprez et al., 1996; Wilson and Tucker, 2004). In zebrafish, Bmp4
misexpression or reduction of the Bmp antagonist Gremlin2 disrupts jaw joint formation, with
Edn1 and Jagged-Notch signalling functioning together to confine grem2 expression to nascent
facial joints (Zuniga et al., 2011). Bmp signalling restricts the jaw joint-forming domain in part
through induction of Hand2, which prevents expansion of nkx3.2 expression into the lower jaw
(Miller et al., 2003). In humans, mutations in EDN1 or its signaling components such as PLCB4
and GNAI3 similarly result in abnormalities of the jaw joint in Auriculo-Condylar Syndrome
(Gordon et al., 2013a; Gordon et al., 2013b; Kido et al., 2013; Leoni et al., 2016; Rieder et al.,
2012). Absence of the jaw articulation is also seen in mice deleted for the Edn1 target gene Dlx5
and Dlx2 (Depew et al., 2005). Somewhat differently, loss of Foxc1 results in pronounced bony
fusions of the upper and lower jaw, a condition known as syngnathia (Inman et al., 2013). In
addition to Edn1, Bmp, and Fgf signaling, Wnt and Hh signaling also play important roles in
14
establishing joint domains (Koyama et al., 2007; Luyten et al., 2009; Rockel et al., 2016). In
mice, Wnt9a/Wnt14 is both necessary and sufficient to specify joint-forming domains in the limb
(Guo et al., 2004; Hartmann and Tabin, 2001), and Ihh is required for limb and jaw joint
development (Koyama et al., 2007; Shibukawa et al., 2007). Conversely, Hh activation has been
found to inhibit joint development through repression of Wnt signaling and Fgf18 expression
(Rockel et al., 2016). Going forward, it will be important to discern the extent to which these
pathways establish early gene expression domains that prefigure joints versus more directly
controlling chondrocyte and other cell fate at the joint.
1.3 Controlling Chondrocyte Identity at the Joint
As the pre-cartilage condensation forms, cells destined for both endochondral bone and
the joint express a number of genes typical of mesenchymal progenitors, including Sox9 and low
levels of Col2a1 (Pitsillides and Ashhurst, 2008). In the anlagen of the endochondral bones,
chondrocytes then upregulate cartilage matrix genes (e.g. Col2a1, Matrilin1, and Aggrecan) and
stratify into proliferative, pre-hypertrophic, and hypertrophic zones (Mackie et al., 2008).
Hypertrophic chondrocytes express Col10a1 and low levels of genes in common with osteoblasts
(e.g. Runx2 and Osterix), mineralize, and undergo apoptosis or transdifferentiation into long-
lived osteocytes (Mayne et al., 1976; von der Mark and von der Mark, 1977; Yang et al., 2014;
Zhou et al., 2014). In contrast, cells within the nascent interzone produce signalling molecules
that help organize the joint and prevent further chondrocyte maturation. An ongoing debate is the
extent to which joint chondrocytes arise from a separate lineage from those of endochondral
bone. Lineage tracing studies based on Col2a1 and Dcx transgenes suggest that endochondral
and joint chondrocytes arise from a common early field of cells in mice (Hyde et al., 2008;
15
Zhang et al., 2011), and the observation that joint chondrocytes enter the endochondral program
in certain zebrafish and mouse mutants would suggest that they share a common, albeit latent,
potential with other chondrocytes in the condensation (Askary et al., 2015; Ray et al., 2015). On
the other hand, there are reports of migratory cells from outside the interzone contributing to
articular chondrocytes and the meniscus in mouse, which would imply an additional separate
origin for at least some joint tissues (Hyde et al., 2008; Pacifici et al., 2006; Ray et al., 2015).
Notwithstanding, it is clear that a number of signalling pathways and transcriptional repressors
converge to potently inhibit chondrocyte maturation at the developing joint surface.
Complex regulation of Bmp signalling at and around the joint suggests a central role of
this pathway in joint progenitor specification (Salazar et al., 2016). A variety of Bmp ligands and
antagonists are expressed in or around the developing interzone in mouse and zebrafish,
including Bmp2, Bmp4, and Bmp7 (Hogan, 1996), Chordin (Archer et al., 2003; Miller et al.,
2003), Noggin (Brunet et al., 1998; Stafford et al., 2011), and Gremlin2 (Zuniga et al., 2011),
and genetic evidence suggests that inhibiting Bmp signalling is critical for joint development.
Loss of Noggin in the Col2a1-expressing lineage of mice disrupts interphalangeal joints (Ray et
al., 2015). On the other hand, loss of Bmp2 and Bmp4 in limb mesenchyme blocks formation of
the elbow joint (Bandyopadhyay et al., 2006). Of particular interest, the Bmp ligands Gdf5,
Gdf6, and Gdf7 are prominently expressed within the joint interzone (Bruneau et al., 1997;
Miller et al., 2003; Reed and Mortlock, 2010; Settle et al., 2003; Storm and Kingsley, 1999;
Wolfman et al., 1997), with Gdf5-Cre activity extensively labelling joint tissues in mice
(Koyama et al., 2008). Gdf5 was first identified as the gene mutated in the brachypodism mouse
(Storm et al., 1994), which displays limb and mild joint defects; mice doubly mutant for Gdf5
and Gdf6 show more extensive joint defects (Settle et al., 2003; Storm et al., 1994; Storm and
Kingsley, 1999). Gdf5 activity is not, however, sufficient to induce joints, as Gdf5 misexpression
16
promotes rather than inhibits cartilage maturation (Coleman and Tuan, 2003; Francis-West et al.,
1999). One possibility is that high Bmp signalling further away from the joint promotes
endochondral ossification, lower Gdf signalling near the joint promotes deeper articular
chondrocyte fates, and a relative absence of Bmp/Gdf signalling at the joint surface preserves
progenitors in an early chondrocyte/mesenchymal state (Figure 1.5). Members of the Tgf
signalling family are also critical for joint morphogenesis (Spagnoli et al., 2007). The Tgfbr2
receptor is expressed in the joint interzone, with its loss in mice resulting in a near complete
absence of the interphalangeal joints due to ectopic hypertrophy of articular chondrocytes (Baffi
et al., 2004; Longobardi et al., 2012; Seo and Serra, 2007). While differentiation of chondrocytes
from human embryonic stem cells in the presence of BMP4 results in a hypertrophic identity,
addition of Tgf maintains chondrocytes in a non-hypertrophic state (Craft et al., 2015; Wu et
al., 2013). Bmp/Gdf and Tgf signalling are thought to signal through a mixture of shared and
distinct SMAD effectors (Salazar et al., 2016). Whereas competition for the common Smad4
effector is predicted to result in cross-inhibitory actions of these pathways, the finding that mice
lacking Smad4 in the Col2a1 lineage have only mild joint defects indicates that the balance of
Bmp/Gdf and Tgf pathways may be more important than the absolute levels of either (Zhang et
al., 2005).
17
The levels, duration, and/or type of Bmp signaling help to specify the different types of chondrocytes at
and around joints: hypertrophic, radial, transitional, and superficial. The Bmp ligands Gdf5/6/7 and
antagonists Chordin (Chd), Noggin (Nog), and Gremlin2 (Grem2) are expressed in the developing
interzone, and Bmp2/4/7 have been reported to be expressed either at a distance from the joint or at the
joint itself. Variable diffusion of these ligands and antagonists may establish different levels of Bmp
signaling, and signaling through Bmpr1a and Bmpr1b receptors could also influence joint fate. Tgfβ
signaling also has an important role to specify joint fates, with the Tgfbr2 receptor enriched at developing
joints.
Figure 1. 5. Critical thresholds of Bmp and Tgfβ signaling in joint development.
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Figure 1. 6. Chondrocyte fate decisions.
(A) Section of the knee joint from an 8-week-old mouse shows that Col2a1 expression (green) is stronger
in growth plate chondrocytes (arrowhead) compared to articular chondrocytes (arrow). Superficial joint
chondrocytes are labelled by treatment of Prg4-CreER; Rosa26:memTomato/memGFP mice with
Tamoxifen three weeks earlier (anti-GFP antibody staining detects Prg4-CreER-converted cells in red).
(B) Compared to the complex mammalian knee joint, the hyoid joint of 6-day-old zebrafish provides a
simplified model for understanding the specification of joint chondrocyte fate. In this example, transient
chondrocytes express both a sox10:dsRed transgene (red) and a col2a1a:GFP transgene (green). In
contrast, joint chondrocytes express sox10 (a member of the SoxE family like Sox9) but much lower
19
levels of col2a1a, suggesting they are immature. (C) Cells within a mesenchymal condensation initially
express Barx1 and then go on to express Sox9, Dcx, and low levels of Col2a1 (in particular an A splice
isoform). In the growth plate, these cells mature into pre-hypertrophic chondrocytes that express high
levels of Col2a1 and Matn1 and then hypertrophic chondrocytes that express Col10a1, Runx2, and other
genes associated with mineralization. In contrast, interzone cells differentiate into articular chondrocytes
that maintain low Col2a1 and instead express Gdf5 and later Prg4. Whereas Wnt9a and Fgf18 promote a
joint fate, Ihh signaling controls cartilage maturation. Specification of articular chondrocyte fate is
promoted through inhibition of cartilage maturation by a number of transcription factors and chromatin
remodellers, including Iroquois proteins (discussed in Chapter 2), Trps1, Nkx3.2, Cux1, Erg, Lrf, and
Hdac1. (D) Under the control of signalling factors including GDF5/6, Wnt9a, Fgf18, TGF β and Noggin,
chondroprogenitor cells mature to form the different chondrocyte layers of the growth plate (darker blue)
or remain relatively immature within the joint (orange cells in light blue matrix). Factors including Trps1,
Has, and Ihh then drive interzone cells to cavitate as superficial zone articular chondrocytes (brown)
begin to produce lubricating molecules such as Prg4.
Whereas lower levels of Bmp signalling are important for specifying joints, less is known
about how such signaling is interpreted in the nucleus to repress the hypertrophic maturation of
articular chondrocytes (Figure 1.6). Cux1 (Lizarraga et al., 2002) and the C-1-1 isoform of the
Ets factor Erg (Iwamoto et al., 2000) are both expressed at developing murine joints and can
repress cartilage differentiation upon misexpression. Loss of Erg in Gdf5+ joint precursors did
not, however, alter joint specification in mice, although it did lead to increased susceptibility to
age- and injury-induced osteoarthritis of the knee (Ohta et al., 2015). As we discuss in the next
chapter, Iroquois family of transcriptional repressors also play a role in joint specification by
inhibit chondrocyte maturation. In addition to Iroquois genes, the Tricho-rhino-phalangeal
20
syndrome gene Trps1 is expressed at joints in zebrafish (Talbot et al., 2010), as well as in early
chondrocytes of the murine growth plate (Kunath et al., 2002; Singh et al., 2016). Loss of Trps1
results in precocious chondrocyte maturation in the condylar cartilage of the murine jaw joint
(Michikami et al., 2012), and cone-shaped epiphyses, joint hypermobility, and osteoarthritis-like
changes to articular cartilage in humans (Maas et al., 2015). Interestingly, IRX5 and TRPS1
proteins form a complex during early neural crest development (Bonnard et al., 2012). Given
their similar expression and requirement in early chondrocytes, it will be interesting to test
whether a similar protein-protein complex functions to inhibit cartilage maturation at joints.
It is becoming increasingly appreciated that covalent modifications of DNA and their
associated histones can have long-term consequences on gene expression. A salient feature of
joint cartilage is that it must be maintained as permanent hyaline cartilage for the life of the
animal. An attractive model then is that epigenetic silencing of the chondrocyte hypertrophy
program helps lock articular cells into a permanent cartilage identity (Furumatsu and Ozaki,
2010). In cultured human chondrocytes, expression of Col10a1 involves active DNA
demethylation of CpG islands in its promoter (Zimmermann et al., 2008). In mice, loss of
Histone Deacetylase 4 (Hdac4) results in accelerated hypertrophy and mineralization of
cartilage, in part through increased histone acetylation and expression of Runx2 (Vega et al.,
2004). Further, treatment of synovium-derived cells from pig with Tgf 1 induces
chondrogenesis, with co-expression of Hdac1 blocking the hypertrophy of these chondrocytes
(Pei et al., 2009). Similarly, the joint-promoting factor Nkx3.2 has been shown to function as a
transcriptional repressor in complex with Hdac1 and the common Bmp/Tgf effector, Smad4,
although the targets of Nkx3.2 remain unclear (Kim and Lassar, 2003). The transcriptional
repressor Lrf has also been shown to act together with Hdac1 to repress chondrogenic
21
differentiation in rodents (Liu et al., 2004). These data suggest that DNA methylation and/or
histone deacetylation of Col10a1, Runx2, and likely other genes associated with chondrocyte
hypertrophy may be important for maintaining permanent cartilage at joints.
1.4 Generation of Synovial Specializations
In addition to generating permanent articular cartilage, cells within the developing joint
express a suite of genes involved in the creation of joint-specific structures (Figure 1.7). A
prominent morphological event in synovial joints is the formation of a cavity between articular
cartilage layers, which is accompanied by the loss of cell-cell contacts and production of
molecular lubricants that promote joint function. In some joints, tissues with features in common
with the articular cartilage layers can exist as menisci that project into the cavity (e.g. in the knee
joint) or as discs between the articular surfaces (e.g. in the mammalian jaw joint). Supportive
ligaments also attach to specific sites in the joint through a transitional structure, the enthesis.
Joints are clearly complex organs that vary greatly depending on their location in the body, with
their development relying on integration of common joint-promoting programs with unique
positional specifiers.
22
Figure 1. 7. Synovial specializations.
Distinct patterns of cavitation of the interzone (green) can generate simple synovial joints (A) or joints
with specialized structures such as menisci (B) and articular discs (C). The synovium (orange) shares
many properties with the fibroblasts ensheathing the menisci and disc (purple). Bony processes (red) act
as attachment points for ligaments (green), with their connection point, or enthesis, consisting of a
transitional cell type.
Cavitation of the developing joint separates the two articular surfaces, yet the cellular
mechanisms underlying this morphogenetic event remain unresolved. Different joints fail to
cavitate in a number of mouse mutants, including the temporomandibular joint in Trps1 mutants
(Michikami et al., 2012), the digit joints in Ihh mutants (Koyama et al., 2007), and elbow and
wrist joints in Osr1; Osr2 double mutants (Gao et al., 2011). In many of these cases it remains
unclear the extent to which lack of cavitation is secondary to earlier defects in joint specification.
In one study, conditional deletion of the Hh receptor Smo in Sox9+ chondroprogenitors resulted
in a failure of the disc to separate from the mandibular condyle but did not affect earlier joint
specification (Purcell et al., 2009). Correlative evidence suggests that cavitation does not require
23
extensive cell death within the developing interzone (Kavanagh et al., 2002). Instead,
asymmetric synthesis of hyularonan (HA), a major component of synovial fluid, may facilitate a
decrease in cell-cell contacts that allows tissue separation (Craig et al., 1990). In the mouse limb,
conditional inactivation of Has2, a member of the HA synthase family, using a Prrx1-Cre line
that is active in early limb mesenchyme, inhibits cavitation of digit joints and leads to an
accumulation of mesenchymal cells in the presumptive joint space (Matsumoto et al., 2009). This
effect is specific to Has2, as deletion of Has1 and/or Has3 does not affect cavitation (Camenisch
et al., 2000).
Synovial joints are also unique in producing lubricating fluid that reduces friction and
protects joint integrity. A critical lubricating protein is the proteoglycan Lubricin, encoded by the
Prg4 gene, which complexes with Aggrecan and HA in the synovial fluid (Jay and Waller,
2014). Within developing joints, Prg4 expression appears only after Gdf5 expression begins to
subside, roughly correlating with initiation of cavitation (Rhee et al., 2005). In addition to
expression in the most superficial cells of articular cartilage, Prg4 expression can also be found
in the synovium, meniscus, and ligaments of mammalian joints. Such shared expression may
reflect combinatorial secretion of Lubricin from all cavity-lining tissues, as well as additional
roles for Lubricin in the lubrication of non-joint tissues such as ligaments (Taguchi et al., 2008).
Although dispensable for joint cavitation, Lubricin is required for the maintenance of many types
of joints with osteoarthritic phenotypes seen in human camptodactyly-arthropathy-coxa vara-
pericarditis (CACP) syndrome patients (Marcelino et al., 1999), mice lacking Prg4 (Koyama et
al., 2014; Rhee et al., 2005). The factors that induce Prg4 expression during later joint
development remain unclear, although a recent study suggests a potential role for mechanical
forces (Ogawa et al., 2014).
24
Menisci and discs are similar to the articular surface in having Prg4-expressing flattened
cells overlying rounder chondrocyte-like cells. These structures arise within the Gdf5-expressing
interzone, with their complex structures influenced by varied patterns of cavitation (e.g. discs are
likely generated by two separate and parallel cavitation events). There are also specialized
endochondral bones at joints that act as ligament attachment sites. These endochondral bones
often form as secondary cartilage outgrowths from the bone surface (e.g. fossa of the mammalian
jaw joint) (Shibukawa et al., 2007). In zebrafish with reduced nkx3.2 function, defects in
articular cartilage are accompanied by loss of joint-associated endochondral bones at the jaw
joint, suggesting a high level of coordination of articular cartilage specification and joint-
associated bone features (Miller et al., 2003). The enthesis is a specialized transitional tissue
between the endochondral bone and ligament at the joint. A common feature of ligament,
enthesis, bone, and cartilage progenitors is expression of Sox9 (Ono et al., 2014; Sugimoto et al.,
2013). It is possible that there exists a common Sox9+ progenitor pool within the early interzone,
with local induction of lineage-specific factors (e.g. Scleraxis for ligaments) resulting in precise
arrays of cartilage, bone, and ligament cells within the joint organ. It remains unclear whether
interzone cells constitute a homogenous cell population that later diverges into distinct lineages,
or whether interzone cells are heterogenous and lineage-biased from initial stages.
1.5 Conclusion
The joint is a complex organ, composed of chondrocytes of different types and an array
of associated connective tissues. Some of these connective tissues, in particular the synovium,
may serve to not only encapsulate the joint cavity but also to act as a source of progenitors for
new chondrocytes and joint tissues. A growing body of work is revealing how Wnt, Bmp/Gdf,
25
Tgfβ, Ihh, Fgf, and other pathways are integrated in the developing joint interzone to control the
expression of transcription factors and chromatin remodellers, which in turn locally specify cell
fates. An important unanswered question is how members of the Tgfβ superfamily are utilized to
specify the different permanent cartilage zones at the articular surface versus the neighboring
transient cartilage that converts into bone. Do the duration and/or relative levels of Bmp versus
Tgfβ signalling drive different cartilage types? Or does signaling through distinct types of
receptors and downstream effectors regulate different fates? A related question is how Gdf and
other signaling pathways regulate non-chondrocyte fates at joints. Chondrocytes, osteoblasts, and
ligament cells all passage through a Gdf5+ interzone cell, with progenitors for each lineage also
expressing Sox9. When do these lineages diverge from each other, and how are the major
signaling pathways integrated differently to achieve these fates? An emerging theme is that there
exists a continuum of cell fates throughout the joint, such as ligament cells that grade into
osteoblasts in the enthesis, superficial chondrocytes that grade into synovial fibroblasts, and
chondrocytes that grade into osteoblasts at the tidemark. Such graded fates may help link
together the different components of the joint into a seamless organ.
A promising future research direction is to understand the extent to which the pathways
that establish an articular cartilage fate function to maintain this fate, which in humans can be for
over one hundred years. An attractive but relatively unexplored possibility is that developmental
pathways establish repressive DNA methylation and chromatin around genes associated with
chondrocyte hypertrophy, thus locking articular chondrocytes into a permanent cartilage
phenotype. Does the postnatal expression of transcriptional repressors such as Iroquois proteins
and Trps1 indicate a requirement for continued reinforcement of silencing of chondrocyte
hypertrophy at the joint surface? And how would injury and decreased lubrication (e.g. due to
PRG4 loss) result in an override of epigenetic repression of chondrocyte hypertrophy? Relatedly,
26
does failure to fully lock articular chondrocytes into a permanent cartilage identity during
development predispose to later osteoarthritis? In the future, a better understanding of how the
different cell types of the joint are specified during development, then maintained and replaced
as needed in the adult, will be critical for guiding new approaches towards both preventing and
reversing osteoarthritis.
1.6 Credits
This literature review has been done in collaboration with Joanna Smeeton and Gage
Crump. The complete work is published as an Advanced Review article entitled “Building and
maintaining joints by exquisite local control of cell fate” in Wiley Interdisciplinary Reviews:
Developmental Biology.
27
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Chapter 2
Iroquois Proteins Promote Skeletal Joint Formation by
Maintaining Chondrocytes in an Immature State
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Figure 2. 1. Regulation of irx7 and irx5a expression and character of the hyoid joint.
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Figure 2. 2. Requirements of Irx7 and Irx5a in hyoid joint formation.
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Figure 2. 3. Irx7 represses chondrocyte maturation.
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Figure 2. 4. Irx genes directly
repress a col2a1a enhancer.
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Figure 2. 5. Progressive restriction of irx7:GFP to the hyoid joint and Hand2-independent inhibition of
irx7 expression by Bmp signaling.
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Figure 2. 6. Hyoid joint and symplectic cartilage defects in independent null alleles for irx7 and irx5a.
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Figure 2. 7. Potential evolutionary history of vertebrate Iroquois genes.
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Figure 2. 8. Potential roles of Irx genes at multiple joints.
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Chapter 3
Unbiased search for novel regulators of joint specification in
zebrafish facial skeleton
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3.1 Abstract
Patterning of the craniofacial skeleton and specification of skeletal elements within it
have been the subject of numerous studies. The majority of these studies have utilized a
candidate based approach to identify major signalling pathways that control facial patterning, as
well as several transcription factors that act in response to those signals. This provides us with a
useful framework for understanding the underlying genetics of craniofacial development.
However, as a result of the inherent limitation of candidate-based approach, some important
factors may have been missed by those studies. Here we take a genomic approach to find novel
regulators of facial patterning and skeletal specification in an unbiased way. Since the facial
joints are made by the neural crest cells in the intermediate domain of the pharyngeal arches, our
main goal is to find genes that are enriched in this domain. To attain this, we use transgenic
zebrafish lines to isolate cells from different domains of the pharyngeal arches, which we then
use for RNA sequencing. We isolated arch cells in this way from wildtype animals as well as
those with perturbed Endothelin1 (Edn1) signalling. Analysis of the gene expression levels in
different domains of the pharyngeal arches guides us towards genes that are differentially
expressed among these domains. This gene list was then refined further by taking into
consideration the known role of Edn1 signalling in promoting the expression of intermediate
genes. Our study identifies a number of genes that are enriched in the intermediate domain of the
arch whose roles have not been investigated in this context. Functional studies on these genes are
likely to provide further insight into patterning of facial skeleton and specification of the joints.
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3.2 Introduction
The facial skeleton is made by differentiating neural crest cells (NCCs) that migrate from
the dorsal tip of the neural tube and populate the pharyngeal arches (Schilling, 1997). Before
skeletal differentiation, NCCs are patterned along the dorsal-ventral axis of the arch into at least
three distinct domains: Dorsal, Intermediate, and Ventral (Alexander et al., 2011). This early
patterning reflects the morphological and cellular differences that later arise among cartilage
elements in different regions of the face. In the zebrafish, NCCs in the dorsal domain of the
pharyngeal arches make flat dorsal facial cartilages, such as palatoquadrate and hyomandibular,
whereas ventral domain cells give rise to rod-shaped ventral cartilages, such as Meckel’s and the
ceratohyal. Joints between the dorsal and ventral parts of the face, such as jaw joint and hyoid
joint, are made by cells in the intermediate domain (Medeiros and Crump, 2012; Mork and
Crump, 2015). Identifying the molecular differences between pharyngeal arch domains is key to
unravelling the genetic programs that specify various skeletal elements of the face.
The major signaling pathways involved in dorsoventral patterning of the face are Jagged-
Notch in the dorsal domain, Endothelin1 (Edn1) in the intermediate and ventral domains, and
Bmp signaling in the ventral domain (Barske et al., 2016). Each of these signals promotes
expression of their target genes specifically in the afore-mentioned domains, and their
perturbation results in mispatterning of the facial skeleton. Loss of Bmp signaling results in loss
of ventral facial structures in both zebrafish and mice (Alexander et al., 2011; Zuniga et al.,
2011). Similarly, mutations affecting Edn1 signaling or its downstream targets, such as Dlx5 and
Dlx6, cause homeotic transformation and/or loss of ventral and intermediate skeletal elements
(Depew et al., 2002; Kurihara et al., 1994; Miller et al., 2000). Moreover, mutation in zebrafish
jag1b, as well as a combined loss of notch2 and notch3 genes, result in malformation of dorsal
66
facial cartilages (Barske et al., 2016; Zuniga et al., 2010). Genetic studies have revealed that
these signaling pathways mutually antagonize each other (Figure 3.1). Edn1 signaling functions
in part through limiting Notch signaling to the dorsal domain, while Notch signaling prevents
dorsal expansion of Edn1 target genes (Barske et al., 2016; Zuniga et al., 2010). Similarly, Bmp
signaling is inhibited in the intermediate domain by downstream targets of Edn1 signaling, such
as grem2b (Zuniga et al., 2011). This three-way antagonism establishes, and refines the borders
of, three domains along dorsoventral axis of the pharyngeal arches.
First and second pharyngeal arches of zebrafish embryo, at 36 hours post fertilization (hpf), and the
skeletal elements that they form in the larval face, at 5 days post fertilization (dpf), are schematized.
Three domains along DV axis are color coded, the same as the cartilage elements that they give rise to.
Edn1 promotes expression of intermediate genes while it inhibits expansion of dorsal and ventral program
into the intermediate domain. On the other hand, Jag1b signaling in the dorsal domain and Bmp signaling
in the ventral domain prevent ectopic expansion of intermediate genes. A, anterior; ch, ceratohyal; D,
dorsal; hm, hyomandibular; M, Meckle’s cartilage; P, posterior; pq, palatoquadrate.
Figure 3. 1. Bmp, Edn1, and Jagged-Notch signaling pattern the facial skeleton of zebrafish.
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The candidate-based genetic studies, such as those mentioned above, have provided a
relatively detailed understanding of how Bmp, Edn1, Jagged-Notch, and their targets interact
with each other in the context of craniofacial development. In contrast, genome wide and
unbiased characterization of the molecular basis of pharyngeal arch patterning has been limited
to a few attempts (Bonilla-Claudio et al., 2012; Jeong et al., 2008). In this study, we performed
genome-wide expression analyses of isolated arch domains to identify their molecular signature.
Our specific goal was to find novel genes enriched in the intermediate domain. Since the
intermediate domain cells give rise to the joints, genes that are specifically expressed in this
domain are likely to have important functions in early specification of the joints and, therefore,
are good candidates for further functional study.
We used two complementary approaches to reliably isolate intermediate specific genes.
First, we performed fluorescence-activated cell sorting (FACS) on dissociated cells from
zebrafish embryos that transgenically label specific cell populations. The transgenic lines that we
used for this purpose were: sox10:dsRed, which labels all the arch NCCs as well as developing
ear; fli1a:eGFP, which labels all the arch NCCs, vasculature, and macrophages; dlx5a:eGFP,
which labels NCCs in the intermediate and ventral domains of the arch as well as parts of the ear;
and hand2:eGFP, which labels the NCCs in the ventral most part of the arch and the heart
(Figure 3.2). By isolating arch NCCs from different domains, and comparing their gene
expression levels, we identified gene sets enriched in each domain.
In the second approach, we compared the transcriptional profile of the arch NCCs
between wildtype animals and animals lacking, or overexpressing, Edn1. Expression of
intermediate specific genes was expected to be reduced in edn1 mutants and increased by Edn1
overexpression, given the role of Edn1 signaling in the intermediate domain. This gave us a
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second criterion by which we evaluated our predicted intermediate genes. Together, our study
provides a number of new intermediate specific genes for secondary analysis.
3.3 Results and Discussion
Isolation of cells from the pharyngeal arch domains
To find genes with spatially restricted expression patterns in the pharyngeal arches, we
needed to isolate cells from different domains and compare their gene expression levels with
each other and with the whole arch population. Arch NCCs, from all domains combined, can be
purified by fluorescence-activated cell sorting (FACS) of dissociated cells from sox10:dsRed;
fli1a:eGFP double transgenic embryos, as described previously (Barske et al., 2016). At 36 hpf,
which is the stage we used for all our experiments, sox10:dsRed labels all arch NCCs (and the
ear), while fli1a:eGFP labels arch NCCs, blood vessels, and macrophages (Figure 3.2). Although
both these transgenes have expression in some cell types outside the pharyngeal arches, they
only overlap in the arch NCC population. This allowed us to isolate exclusively arch NCCs with
minimal contamination of other cell types. At the same time, we collected cells that are double
negative or single positive for just one of the two transgenes. These were later used to exclude
genes with high expression in other tissues from our gene lists. Following the same procedure,
we isolated arch NCCs from edn1
-/-
; sox10:dsRed; fli1a:eGFP embryos, which we used to find
genes that are downregulated in the arch upon loss of Edn1 signaling.
To isolate cells from sub-regions of the arch, we sorted cells doubly positive for
hand2:eGFP; sox10:dsRed, which are the ventral most cells in the arches, and also dlx5a:eGFP;
sox10:dsRed, which are cells of the intermediate and ventral domains of the arch as well as parts
of the developing ear (Figure 3.2). Because hand2:eGFP; sox10:dsRed double positive cells
69
comprise a small fraction of the arch, the number of cells acquired in an individual experiment
was low (~18,000 cells). Therefore, to get a more reliable estimate of the gene expression levels
in the ventral domain, we repeated the sort and sequencing of hand2:eGFP; sox10:dsRed double
positive population twice.
70
(A) Confocal imaging of a 36 hpf fli1a:eGFP; sox10:dsRed embryo shows that the two transgenes
intersect in the arches (yellow), while ear cells are labeled with only dsRed (red) and the vasculature
around the eye expresses only GFP (green). (B) An image of a dlx5a:eGFP; sox10:dsRed embryo, at 36
hpf, shows that the overlap between the two transgenes labels intermediate and ventral domains. (C) 36
hpf hand2:eGFP; sox10:dsRed embryo showing the colocalization of the two transgene in the ventral
domain. D, dorsal; I, intermediate; V, ventral.
Figure 3. 2. Transgenic lines labeling different regions of the pharyngeal arch help isolate specific
populations.
71
Transcriptomic analysis reveals genes that are enriched in the intermediate domain
After RNA sequencing of the above-mentioned cell populations, read counts were
normalized to yield RPKM (Reads Per Kilobase of transcript per Million mapped reads) values
for each sample. Because some genes in some samples didn’t have any reads, a low base
number, 0.0001, was added to all RPKM values to avoid division by zero in the calculation of
fold-changes. We are only interested in the genes that are expressed in the arches, therefore we
excluded genes with RPKM values lower than 0.1 in the wild-type fli1a:eGFP; sox10:dsRed
double positive sample. Moreover, we only included genes that are enriched 1.5 fold or higher in
the wild-type fli1a:eGFP; sox10:dsRed double positive sample (arch NCC) compared to either of
the single positive samples (other cell types, including ear, vasculature, and macrophages). This
leaves 1347 “arch genes”. To enrich this list for genes with intermediate specific expression
pattern, we filtered for genes with RPKM values at least 1.5 times higher within dlx5a:eGFP;
sox10:dsRed double positive versus the fli1a:eGFP; sox10:dsRed double positive cell
populations. There were 293 such genes (Table 2.1 left column). All subsequent analysis used
this list as a starting point.
To narrow this list down, we took two approaches. In the first approach we accounted for
the fact that ventral genes were also included in the list because dlx5a:eGFP also labels the
ventral domain. To exclude ventral genes, we intersected our list with genes that have at least 1.5
times higher RPKM in the dlx5a:eGFP; sox10:dsRed double positive sample compared to both
of hand2:eGFP; sox10:dsRed double positive samples. This shrank our list from 293 to 88 genes.
The second approach was based on the fact that Edn1 signaling promotes the expression of
intermediate genes. Therefore, we expect intermediate genes to be reduced in edn1
-/-
embryos.
So we further refined our list to include only those genes that underwent a 1.5 times or greater
reduction in edn1
-/-
; sox10:dsRed; fli1a:eGFP arch NCCs when compared to the wildtype cell
72
population. This left 39 “intermediate genes” (Table 2.1 middle column). Alternatively, the 293
intermediately enriched genes were intersected with our previously published Edn1 activated
gene list (Barske et al., 2016): this identified 32 “intermediate genes” (Table 2.1 right column).
73
Table 3. 1. List of genes enriched in the intermediate domain of zebrafish pharyngeal arches.
Enriched in dlx5a+; sox10+ vs. fli1a+; sox10+
… AND enriched in
dlx5a+; sox10+ vs.
hand2+; sox10+
AND reduced in edn1
-/-
… AND
activated by
Edn1
ago3b ahdc1 alg12 alyref angptl2b ankrd10a appl2 araf arl16
arl4cb arsk asf1ba barx1 BEND2 (2 of 2) bmp5 bmpr1ba
bmpr1bb btbd2a btr07 BX324179.3 BX548028.1 BX548067.2
BX569782.1 BX571691.1 BX649198.1 C2CD4C (1 of 2)
C9H21orf33 CABZ01033394.1 CABZ01045617.1
CABZ01050663.1 CABZ01054394.4 CABZ01064672.1
CABZ01066717.1 CABZ01066720.1 CABZ01110379.1 cbx1a
cby1 ccdc103 CCNJ cdc14ab cdca4 cep72 chodl chordc1a
chrna2a chst10 chtopa cited2 CKN150339.1H9orf41 cmtm4
cnpy2 cnpy4 COX5B (1 of 3) CR450785.2 CR769782.2 creb5a
crispld1b ctgfb CU104709.1 cx30.9 cyp2ad3 cyp2x7 dact3a
dctpp1 dexi dlx1a dlx2a dlx3b dlx4a dlx4b dlx6a dpcd efcab1
emx2 ETAA1 (2 of 2) fastkd3 fen1 fgf3 fgfbp1b fgfbp2a fgfr2 fibina
figf FKBPL FO704710.3 foxc1b foxd1 foxf1 foxf2a foxk2 fsta
fubp1 fzd8a gdf10a gdf11 gpr161 grb2a grem2b gsc hand2 hapln2
has2 her6 hic1l hmcn2 hnrnpa0b hoxa2b hoxb1a hoxb2a id3 ift22
ift57 igfbp5b il15l iqcb1 irx7 khsrp klhl21 lhx6 lhx8a lhx8b lin28a
LIPC (2 of 2) lix1l lrig1 lrrc8c LRRN3 mab21l2 map1lc3c map9
MDFIC med1 med14 meis1a meis2a mettl14 mier1a mrrf
ms4a17a.11 ms4a17a.7 ms4a17a.8 MSANTD1 msxe mtf2 mycn
nog2 notch2 nr2f1b nrn1b nucks1b nxf1 ociad2 ompa p2ry11 pax9
PGBD1 phf12b phlda3 pitx1 pkd2 pmp22a ppp1r3ca ptch1 pygo2
rab34a rbbp8 rbm18 rcn2 rfc1 rgmd rhot1b rnmtl1a rpap2 rpe
rsad2 rtn4b sall4 samd1b satb2 sema3b senp3b sephs1 sept5a
shox shprh si:ch211-107p11.3 si:ch211-108d22.2 si:ch211-
130h14.6 si:ch211-134c9.2 si:ch211-159i8.4 si:ch211-165b19.11
si:ch211-196f5.9 si:ch211-197g15.9 si:ch211-209n20.58
si:ch211-222l21.1 si:ch211-238e22.1 si:ch211-262i1.5 si:ch211-
283e2.7 si:ch211-39k3.2 si:ch211-66k16.27 si:ch73-110p20.1
si:ch73-14h10.2 si:ch73-166c6.1 si:ch73-281n10.2 si:ch73-
341p18.7 si:ch73-351f10.4 si:ch73-386o14.1 si:dkey-121h17.7
si:dkey-125i20.2 si:dkey-179o14.4 si:dkey-182i3.10 si:dkey-
184n3.2 si:dkey-19d21.2 si:dkey-207l24.2 si:dkey-20i10.5 si:dkey-
244a7.1 si:dkey-245m3.2 si:dkey-253d23.9 si:dkey-269o24.7
si:dkey-270b7.4 si:dkey-286h21.1 si:dkey-34o23.2 si:dkey-
43b14.9 si:dkey-57c15.9 si:dkey-60d5.3 si:dkey-90a13.10 si:dkey-
93m18.6 si:dkey-96n2.1 si:dkeyp-106c3.2 si:dkeyp-10a3.2
si:dkeyp-38g8.5 si:dkeyp-44d3.4 si:dkeyp-53e4.1 si:dkeyp-86b9.5
si:rp71-1c10.10 slc16a12b SLC16A6 (3 of 3) slc38a3b slc38a4
slc48a1b smad6a smarcc2 smo snx10a sox11a sox9a spon2b
srsf10a ssbp4 stmn1a stom styxl1 tbx22 tctn1 tdp1 tead1b tead3a
thoc7 tmed5 tmem107l tmem119b tmem17 tmem231 tnfsf12
trmt10b tspan7 ttc26 twist1a twist1b txlng vasna vegfaa wrnip1
wu:fe11b02 yap1 ylpm1 zbtb49 zfp64 zgc:101130 zgc:101744
zgc:113210 zgc:113336 zgc:113413 zgc:114037 zgc:153154
zgc:153935 zgc:162612 zgc:162936 zgc:165555 zgc:194469
zgc:92664 znf384l znf644a
bmpr1bb
CABZ01033394.1
chrna2a
ctgfb
dlx3b
dlx4a
dlx4b
dlx6a
fgfbp1b
fibina
foxc1b
fsta
hapln2
igfbp5b
irx7
ms4a17a.7
ms4a17a.8
nrn1b
ompa
rsad2
si:ch211-130h14.6
si:ch211-165b19.11
si:ch211-222l21.1
si:ch211-283e2.7
si:ch73-351f10.4
si:dkey-184n3.2
si:dkey-20i10.5
si:dkey-269o24.7
si:dkey-90a13.10
si:dkeyp-106c3.2
si:dkeyp-86b9.5
slc16a12b
spon2b
tbx22
tmem17
tnfsf12
wu:fe11b02
zgc:114037
zgc:165555
ankrd10a
arl4cb
bmpr1ba
cby1
ctgfb
CU104709.1
dlx3b
dlx4a
dlx4b
dlx6a
figf
foxc1b
fsta
gpr161
grem2b
lhx8b
lrig1
lrrc8c
ms4a17a.11
ms4a17a.7
msxe
mycn
nog2
notch2
phlda3
ptch1
sox9a
spon2b
stom
tbx22
tmem17
zgc:194469
74
The predicted intermediate gene lists include previously characterized genes as well as novel
candidates
As expected, our lists of intermediate genes include many of the genes that have been
reported to be expressed in the intermediate domain of the pharyngeal arches. This includes
dlx4a, dlx6a, dlx3b, and dlx4b; targets of Edn1 signaling, important for setting up an
intermediate identity (Talbot et al., 2010). irx7, which promotes development of the hyoid joint
as discussed in chapter 2 (Askary et al., 2015), and grem2b and bmpr1bb, that are involved in the
regulation of Bmp signaling in the intermediate domain (Alexander et al., 2014; Zuniga et al.,
2011). Moreover, expression and function of tbx22 (Swartz et al., 2011), mycn (Alexander et al.,
2014), and notch2 (Zuniga et al., 2010) have been addressed before in the context of zebrafish
craniofacial development. Apart from the previously studied intermediate genes, our lists include
two other types of genes: genes known to be expressed in the intermediate domain but whose
role in patterning and specification of zebrafish craniofacial skeleton is not understood (e.g. fsta,
ctgfb, foxc1b, igfbp5b, and lrig1), and genes not previously known to be expressed in the arch
(fgfbp1b and spon2b). Figure 3.3 shows the expression pattern of some of these genes revealed
by in situ hybridization. Both of these categories offer exciting candidates to be studied further in
depth in future.
3.4 Conclusion
As we showed in the previous chapter, the facial skeleton of zebrafish offers a tractable
and relatively simple model for the study of joint development. Through a global analysis of
gene expression, we identified a number of genes specifically expressed in the joint forming
75
domain of the pharyngeal arch. Their intermediate specific expression pattern suggests roles in
establishing the intermediate identity, specifying the joints, or regulating the differentiation of
chondrocytes. Some genes, such as fsta and nog2, are implicated in regulation of Bmp signaling
in other contexts (Dalcq et al., 2012; Furthauer et al., 1999). As we showed in chapter 2, Bmp
signaling is an essential step in joint development. In future, it will be interesting to verify the
expression pattern of all our predicted intermediate genes by RNA in situ hybridization.
Subsequently, the role of the verified candidates in developing joints can be examined through
loss- and gain-of-function studies.
Figure 3. 3. Intermediate specific expression of the candidate genes can be verified by RNA in situ
hybridization.
(A) at 36hpf, fsta (red) is expressed in the intermediate domain of the second arch (arrow) and in
pharyngeal pouch endoderm (asterisks). Immunostaining for GFP (green) in sox10:GFP-CAAX
transgenic embryos is used to show the outline of the arches. (B) ctgfb (red) expression in the pharyngeal
arches is also restricted to the intermediate domain (arrow). (C,D) colorimetric in situ hybridization for
76
igfbp5b (C) and lrig1 (D) at ~48hpf shows that their expression pattern in the arches. Colorimetric in situ
images courtesy of ZFIN (Thisse, 2001; Thisse, 2005).
3.5 Materials and Methods
Fish husbandry and zebrafish lines
The Institutional Animal Care and Use Committees of the University of Southern California
approved all zebrafish (Danio rerio) procedures used in this study. We used the following
previously reported zebrafish lines for this study: Tg(sox10:dsRED)
el10
(Das and Crump, 2012),
Tg(fli1a:EGFP)
y1
(Lawson and Weinstein, 2002), Tg(dlx5a:EGFP)
j1073aGt
(Talbot et al., 2010),
Tg(hand2:EGFP)
pd24
(Yin et al., 2010), Tg(sox10:EGFP-CAAX)
el375
(Askary et al., 2015), and
sucker/edn1
tf216
(Miller et al., 2000).
Preparation of FACS-sorted cell populations
To generate doubly transgenic fish lines, sox10:dsRed animals were crossed to
fli1a:eGFP, dlx5a:eGFP, and hand2:eGFP animals. The resulting sox10:dsRed; fli1a:eGFP and
sox10:dsRed; hand2:eGFP animals were then each incrossed to make the embryos for their
corresponding experiments. sox10:dsRed; dlx5a:GFP fish, however, were crossed to
sox10:dsRED animals to avoid homozygosity of dlx5a
j1073aGt
allele. To produce edn1
-/-
;
fli1a:eGFP; sox10:dsRed embryos, edn1
+/-
; fli1a:eGFP; sox10:dsRed animals were incrossed
and then mutant embryos were selected under the fluorescent microscope at approximately 34
hpf based on the reduced distance between the bottom of the first pharyngeal pouch and the
ventral border of the arches. Transgenic embryos were dissociated and prepared for FACS at
approximately 36 hpf as described previously (Barske et al., 2016). A MoFlo Astrios instrument
77
(Beckman-Coulter, Brea, CA, USA) was then used to sort cells based on GFP and dsRed
expression. Cells were collected directly into RLT lysis buffer (Qiagen).
Preparation of cDNA library and RNA sequencing
After the FACS, total RNA was immediately extracted using the RNeasy Micro kit
(Qiagen) following the manufacturer’s instructions. This RNA was then used to make cDNA and
sequencing libraries as described previously (Barske et al., 2016). In brief, the integrity and
quantity of extracted RNA were assessed on a Bioanalyzer Pico RNA chip (Agilent, Santa Clara,
CA). Total RNA was then used to synthesize cDNA using the SMARTer V3 kit (Clontech,
Mountain View, CA), according to the manufacturer’s protocol. The size and amount of the
resulting cDNA were then confirmed by Bioanalyzer. Following sonication of the resulting
cDNA on an S2 ultrasonicator (Covaris, Woburn, MA), sequencing libraries were constructed
using the Kapa Hyper prep kit (Kapa Biosystems,Wilmington, MA) and NextFlex adapters (Bioo
Scientific, Austin, TX). The quality of the libraries was then assessed by Bioanalyzer analysis
and qPCR (Kapa library quantification kit). Sequencing was performed on Illumina HiSeq 2000
(50-bp paired end reads) and NextSeq 500 (75-bp paired end reads) machines (Illumina, San
Diego, CA).
RNA Sequence analysis
Partek flow software was used to align the sequencing reads to zebrafish GRCz10
genome assembly (Ensembl_v80) and quantify the aligned reads as described previously (Barske
et al., 2016). Briefly, a minimum end quality level (Phred) of 20 and a minimum read length of
25 was used to trim both ends of the read. Acceptable reads were then aligned using TopHat 2
78
algorithm and quantified by Partek E/M algorithm. RPKM values were then exported and used in
the downstream analysis. Filtering of the gene lists as well as sorting and intersecting them was
done in MS Excel.
in situ hybridization and embryo imaging
Probes were generated for zebrafish fsta and ctgfb using PCR amplification from cDNA.
Following primers were used in amplification of probe templates: 5’-
GGCTACACCCCATTCTGCTA-3’ and 5’- GGCATCCAGACAACTCGAAA-3’ for ctgfb and
5’-GGAAGACCAGGAGGATGACGATG-3’ and 5’- TCCGTTGACCTTGTGTTCGC-3’ for
fsta. PCR products were then subcloned into pCR-Blunt II TOPO vector (Invitrogen). Following
sequence confirmation, digoxigenin (DIG)-labelled antisense probes were synthesized from
linearized plasmids using T7 or SP6 RNA polymerase (Roche). Fluorescent in situ hybridization
followed by immunostaining for GFP was carried out as previously reported (Askary et al.,
2015). Confocal microscopy was performed on a ZEISS LSM 800 microscope using ZEN
software.
79
3.6 References
Alexander, C., Piloto, S., Le Pabic, P., and Schilling, T.F. (2014). Wnt signaling interacts with bmp and edn1 to
regulate dorsal-ventral patterning and growth of the craniofacial skeleton. PLoS Genet 10, e1004479.
Alexander, C., Zuniga, E., Blitz, I.L., Wada, N., Le Pabic, P., Javidan, Y., Zhang, T., Cho, K.W., Crump, J.G., and
Schilling, T.F. (2011). Combinatorial roles for BMPs and Endothelin 1 in patterning the dorsal-ventral axis of the
craniofacial skeleton. Development 138, 5135-5146.
Askary, A., Mork, L., Paul, S., He, X., Izuhara, A.K., Gopalakrishnan, S., Ichida, J.K., McMahon, A.P., Dabizljevic,
S., Dale, R., et al. (2015). Iroquois Proteins Promote Skeletal Joint Formation by Maintaining Chondrocytes in an
Immature State. Dev Cell 35, 358-365.
Barske, L., Askary, A., Zuniga, E., Balczerski, B., Bump, P., Nichols, J.T., and Crump, J.G. (2016). Competition
between Jagged-Notch and Endothelin1 Signaling Selectively Restricts Cartilage Formation in the Zebrafish Upper
Face. PLoS Genet 12, e1005967.
Bonilla-Claudio, M., Wang, J., Bai, Y., Klysik, E., Selever, J., and Martin, J.F. (2012). Bmp signaling regulates a
dose-dependent transcriptional program to control facial skeletal development. Development 139, 709-719.
Dalcq, J., Pasque, V., Ghaye, A., Larbuisson, A., Motte, P., Martial, J.A., and Muller, M. (2012). RUNX3, EGR1
and SOX9B form a regulatory cascade required to modulate BMP-signaling during cranial cartilage development in
zebrafish. PLoS One 7, e50140.
Das, A., and Crump, J.G. (2012). Bmps and id2a act upstream of Twist1 to restrict ectomesenchyme potential of the
cranial neural crest. PLoS Genet 8, e1002710.
Depew, M.J., Lufkin, T., and Rubenstein, J.L. (2002). Specification of jaw subdivisions by Dlx genes. Science 298,
381-385.
Furthauer, M., Thisse, B., and Thisse, C. (1999). Three different noggin genes antagonize the activity of bone
morphogenetic proteins in the zebrafish embryo. Dev Biol 214, 181-196.
Jeong, J., Li, X., McEvilly, R.J., Rosenfeld, M.G., Lufkin, T., and Rubenstein, J.L. (2008). Dlx genes pattern
mammalian jaw primordium by regulating both lower jaw-specific and upper jaw-specific genetic programs.
Development 135, 2905-2916.
Kurihara, Y., Kurihara, H., Suzuki, H., Kodama, T., Maemura, K., Nagai, R., Oda, H., Kuwaki, T., Cao, W.H.,
Kamada, N., et al. (1994). Elevated blood pressure and craniofacial abnormalities in mice deficient in endothelin-1.
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Lawson, N.D., and Weinstein, B.M. (2002). In vivo imaging of embryonic vascular development using transgenic
zebrafish. Dev Biol 248, 307-318.
Medeiros, D.M., and Crump, J.G. (2012). New perspectives on pharyngeal dorsoventral patterning in development
and evolution of the vertebrate jaw. Dev Biol 371, 121-135.
Miller, C.T., Schilling, T.F., Lee, K., Parker, J., and Kimmel, C.B. (2000). sucker encodes a zebrafish Endothelin-1
required for ventral pharyngeal arch development. Development 127, 3815-3828.
Mork, L., and Crump, G. (2015). Zebrafish Craniofacial Development: A Window into Early Patterning. Current
topics in developmental biology 115, 235-269.
Schilling, T.F. (1997). Genetic analysis of craniofacial development in the vertebrate embryo. BioEssays : news and
reviews in molecular, cellular and developmental biology 19, 459-468.
Swartz, M.E., Sheehan-Rooney, K., Dixon, M.J., and Eberhart, J.K. (2011). Examination of a palatogenic gene
program in zebrafish. Dev Dyn 240, 2204-2220.
Talbot, J.C., Johnson, S.L., and Kimmel, C.B. (2010). hand2 and Dlx genes specify dorsal, intermediate and ventral
domains within zebrafish pharyngeal arches. Development 137, 2507-2517.
Thisse, B., Pflumio, S., Fürthauer, M., Loppin, B., Heyer, V., Degrave, A., Woehl, R., Lux, A., Steffan, T.,
Charbonnier, X.Q. and Thisse, C. (2001). Expression of the zebrafish genome during embryogenesis (NIH R01
RR15402). In ZFIN Direct Data Submission (http://zfinorg).
Thisse, C., and Thisse, B. (2005). High Throughput Expression Analysis of ZF-Models Consortium Clones. (ZFIN
Direct Data Submission (http://zfin.org)).
Yin, C., Kikuchi, K., Hochgreb, T., Poss, K.D., and Stainier, D.Y. (2010). Hand2 regulates extracellular matrix
remodeling essential for gut-looping morphogenesis in zebrafish. Dev Cell 18, 973-984.
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BMP and Endothelin 1 signaling in dorsoventral patterning of the facial skeleton. Development 138, 5147-5156.
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Chapter 4
Ancient origin of lubricated joints in bony vertebrates
81
4.1 Abstract
Synovial joints are lubricated connections between the bones of our body. These joints
are commonly affected in arthritis. It is assumed that synovial joints first evolved as vertebrates
came onto land, with ray-finned fishes lacking lubricated joints. Here we examine the expression
and function of Prg4/Lubricin, a critical lubricating protein in mammalian synovial joints, within
ray-finned fishes. We find that Prg4 homologs are specifically enriched at the jaw and pectoral
fin joints of zebrafish, stickleback, and gar, with genetic deletion of the zebrafish prg4b gene
resulting in the same age-related degeneration of joints as seen in Lubricin-deficient mice and
humans. Our data support lubricated synovial joints evolving much earlier than currently
accepted, at least in the common ancestor of all bony vertebrates. Establishment of the first
arthritis model in the highly regenerative zebrafish will offer unique opportunities to understand
the aetiology and possible treatment of synovial joint disease.
4.2 Introduction
Synovial joints allow for free movement between adjacent bones and are characterized by
a fluid-filled cavity separating layers of hyaline articular cartilage. The synovial cavity is
enclosed by a membrane, which is often strengthened externally by a fibrous capsule and
contains lubricating molecules, such as hyaluronic acid and Lubricin, that reduce friction at the
joint surface (Koyama et al., 2014; Rhee et al., 2005). A prevailing hypothesis is that lubricated
synovial joints first evolved in tetrapods in response to newfound mechanical challenges
imposed on the weight-bearing joints of nascent limbs (van der Kraan, 2013a, b) (Figure 4.1A).
Whereas previous histological studies had suggested that the jaw joints of lungfish (a lobe-finned
fish like humans) (Bemis, 1986), and potentially longnose gar and sturgeon (ray-finned fishes)
82
(Haines, 1942), have synovial-like morphology, the extent to which these joints are molecularly
and functionally similar to tetrapod synovial joints had remained untested. In particular, the
assumption that ray-finned fishes lack the sophisticated types of lubricated joints found in
humans has hampered the use of the zebrafish model for the study of synovial joint diseases such
as arthritis. By examining the expression and function of homologs of a critical lubricating
protein of mammalian joints, Lubricin, we provide evidence that certain joints of adult zebrafish
are indeed true synovial joints.
4.3 Results
Synovial-like morphology of the jaw and fin joints of ray-finned fishes
Given the suggested synovial-like morphology of jaw joints in several fishes, we
examined whether joints of the widely used teleost species, the zebrafish (Danio rerio), also
display synovial morphology (Figure 4.1A). Manual opening and closing of the mouth in fixed
Alcian-Blue-stained animals show that the jaw joint, an articulation between the anguloarticular
and quadrate bones, moves in a single plane, resembling a hinge joint (Figure 4.1B). While study
of zebrafish has contributed to our understanding of the embryonic development of the jaw joint,
zebrafish larvae at the most commonly studied stage, 6 days post fertilization (dpf), show little
evidence of a synovial cavity (Miller et al., 2003). However, whether this joint acquires synovial
characteristics later had not been described. Our histological investigations revealed the variable
presence of a partial cavity as early as 14 dpf, and a prominent and consistently present cavity by
28 dpf (Figure 4.1C,D). In adult zebrafish (12 months post-fertilization, mpf), distinct layers of
flattened articular chondrocytes line the jaw joint cavity, with hypertrophic chondrocytes located
beneath the articular surface (Figure 4.1E,F). We also observed similar cavities lined by flattened
83
chondrocytes in joints of the pectoral fin in adult zebrafish, in particular between the proximal
and distal radials and between the marginal ray and scapula (Figure 4.1B,G). We next used sox10
and tricho-rhino-phalangeal syndrome 1 (trps1) transgenes (Askary et al., 2015) to visualize jaw
joint cavitation over time in living zebrafish. At 3, 7, and 14 dpf, the early chondrocyte marker
trps1:GFP labels the subset of sox10:dsRed+ chondrocytes at the joint surface, with a partial
cavity variably apparent at 14 dpf (Figure 4.2A-C). At 1, 2 and 8 mpf, trps1:GFP is maintained
in articular chondrocytes as the cavity expands, with a subset of cells marked by sox10:dsRed
(Figure 4.2D-F). In the jaw of the relatively small zebrafish, the presence of a cavity and joint-
lining cells, with a minimal fibrous capsule, is consistent with it being a synovial-like joint in
miniature, similar to the homologous incudomalleolar synovial joint of the mammalian middle
ear (Whyte et al., 2002).
To examine whether synovial-like morphology is a conserved feature of ray-finned fish,
we also examined juveniles of the distantly related teleost fish, the three-spined stickleback
(Gasterosteus aculeatus), and an outgroup of teleost fish, the ray-finned spotted gar (Lepisosteus
oculatus) (Braasch et al., 2016). Both stickleback and spotted gar jaw joints displayed synovial
cavities lined by flattened cells (Figure 4.1H,I). In the jaw joint of the larger spotted gar
(standard length 10.2 cm), an internal one-cell-thick membrane and thick external fibrous
capsule enclosed the cavity, with joint cartilage divided into the same superficial, transitional,
radial, and calcified layers seen in mammalian synovial joints (Figure 4.1J,K). The presence of
these additional morphological features in the gar jaw supports synovial-like features being an
ancestral property of bony vertebrates.
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Figure 4. 1. Synovial-like morphology of jaw and fin joints in ray-finned fish.
(A) Phylogenetic tree contrasts the old model of synovial joint evolution (grey asterisk) with the new
model of synovial joints evolving in a common precursor of all bony vertebrates (black asterisk). (B)
Alcian Blue-stained adult zebrafish and accompanying diagrams show the pectoral fin joints, and jaw
joints in open and closed positions. (C-E, G-I) Sections of 14 dpf (n = 4), 1 mpf (n = 3), and adult (n = 6)
zebrafish jaw joints; ray-scapula joint in the adult zebrafish pectoral fin (n = 4); and stickleback (1 mpf, n
= 3) and spotted gar (10.2 cm, n = 3) jaw joints. Sections are stained by H&E (C, D, G-I) or trichrome
(E). Articular chondrocytes (black arrowheads) line the cavity. (F) Schematic of adult jaw joint shows
bone (red), cartilage (blue, lighter shade indicates articular), and synovial membrane (green). (J, K)
Magnifications of (I) show the synovial membrane (arrow), fibrous capsule (asterisk) and multilayered
articular cartilage (K). Scale bar in h, 100 μm; all other panels, 50 μm. aa: anguloarticular; q: quadrate; sc:
85
scapula; r: ray; pr: proximal radial; dr: distal radial; M: Meckel’s; pq: palatoquadrate; s: superficial; t:
transitional; rd: radial layer; c: calcified cartilage; b: bone.
Figure 4. 2. Live imaging of jaw joint cavitation.
(A-F) In double transgenic zebrafish, sox10:DsRed marks all chondrocytes and trps1:GFP labels nascent
joint chondrocytes (arrow) and perichondrial cells at 3 and 7 dpf. By 14 dpf, a partial cavity (asterisk) was
evident in 1/6 animals. In 3/3 animals at 1 mpf, 4/4 at 2 mpf, and 4/4 at 8 mpf, a fully formed cavity is
evident and sox10:DsRed is expressed in a subset of trps1:GFP+ articular chondrocytes. Scale bar, 50μm.
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Expression of Prg4 homologs at joints of diverse fishes
Given the synovial-like morphology of several types of joints in fish, we next examined
whether the chondrocytes lining these joints share a common molecular signature with those of
mammalian joints. Chondrocytes lining mammalian synovial joints differ from those in the
growth plate by expressing Prg4, which encodes a Lubricin proteoglycan that forms a cross-
linked network with hyaluronan and Aggrecan that reduces friction across the joint surface (Jay
and Waller, 2014). Consistent with the appearance of a synovial cavity in juvenile stages, we
find that the joint-lining cells of the zebrafish jaw express prg4b starting from 15 dpf and
continuing throughout adulthood (Figure 4.3A-D). We observed much weaker levels of
expression at other joints of the face, such as the midline ceratohyal-ceratohyal joint (Figure
4.3A, arrowhead) and hyoid joint (Figure 4.3E), which lack synovial morphology. Expression of
prg4b was not detected in the jaw joint at earlier stages (7 dpf, data not shown), consistent with
the late onset of Prg4 expression at mammalian joints (Rhee et al., 2005). In addition, prg4b
expression appeared in joint-lining cells of the synovial-like radial and ray-scapula articulations
of the pectoral fin at 3 mpf, but not in the non-synovial intervertebral discs (Figure 4.3F,G).
Zebrafish also expressed prg4b outside of joints, including conserved expression with
mammalian Prg4 in liver (Ikegawa et al., 2000) and possibly ligaments (Sun et al., 2006) (Figure
4.3A,G). Similar to zebrafish, stickleback expressed prg4b and gar expressed prg4 in joint-lining
cells of the juvenile jaw (Figure 4.3 H-J). In contrast, the related prg4a gene is not enriched at
the jaw joint in either zebrafish or stickleback, instead showing expression throughout cartilage
(Figure 4.4). Of note, the lineage leading to the spotted gar diverged before the teleost genome
duplication (Amores et al., 2011), resulting in zebrafish and stickleback having two Prg4 co-
orthologs and gar a single ortholog (Figure 4.5). Analysis of the single prg4 gene in gar therefore
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reveals that enriched expression of Prg4 within articular chondrocytes existed before the
divergence of ray-finned and lobe-finned vertebrates.
We next examined whether articular chondrocytes of the zebrafish jaw display other
features of chondrocytes lining mammalian synovial joints, including enriched expression of
hyaluronan synthase (Has) enzymes in the radial layer (Hiscock et al., 2000) and lower levels of
types II and X Collagen, Aggrecan, and Matrilin compared to growth plate chondrocytes (Khan
et al., 2007). Consistently, we observed relative depletion of col10a1, acana, and matrilin1
mRNA and Col2a1a and Aggrecan protein in articular versus deeper chondrocytes, as well as
enriched expression of has3 in the radial layer of the juvenile zebrafish jaw joint (Figure 4.3 K-
M, and Figure 4.4). Although none of these markers are exclusive to synovial joints, the shared
expression of Lubricin and hyaluronan synthase enzymes and relative absence of cartilage
maturation genes (e.g. Collagen II/X, Aggrecan, Matrilin1) demonstrates a common molecular
signature between articular chondrocytes of the zebrafish jaw and mammalian synovial joints.
88
89
Figure 4. 3. Expression of Prg4 genes in articular chondrocytes of ray-finned fish.
(A-G) prg4b expression in articular chondrocytes of the zebrafish jaw joint (boxed region in A, B-D),
hyoid joint (E); ray-scapula joint (F); and vertebrae (G). prg4b is also expressed in the liver (asterisk),
possibly in ligaments above the vertebrae, and weakly at the ceratohyal-ceratohyal joint (arrowhead). n =
3 each. (H-J) Expression of stickleback prg4b (1 mpf, n = 3) and gar prg4 (10.2 cm, n = 3) in jaw joint
articular chondrocytes (J, magnification of I). (K-M) Exclusion of acana, col10a1, and matn1 expression
from articular chondrocytes of the zebrafish jaw. n = 3 each. Scale bar, 50μm. ih: interhyal. See also
Figure 4.4 and Figure 4.5.
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Figure 4. 4. Gene expression within the zebrafish and stickleback jaw joints.
(A-D) In situ hybridization reveals broad chondrocyte expression of prg4a but no enrichment within jaw
joint articular chondrocytes (C-D, magnified views). (E) Zebrafish, has3 is expressed in chondrocytes just
underneath the articular surface (arrow) and in a small number of cells within the growth plate
(arrowhead). (F) Expression of matn1 is excluded from superficial chondrocytes of the zebrafish jaw joint
at 1 mpf. (G, H) Immunofluorescence staining for Col2a1 and Aggrecan protein reveals broad cartilage
expression yet exclusion from jaw articular chondrocytes. n = 3 each panel. Scale bars, 50 μm.
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(A) Maximum Likelihood phylogeny of vertebrate Prg4 proteins generated using the LG model in
PhyML (Guindon et al., 2009). The tree was rooted with spotted gar and human Vitronectin (Vtn)
proteins. In the ray-finned lineage, after divergence of the holostean lineage (spotted gar and bowfin), the
teleost genome duplication (TGD) generated Prg4a and Prg4b found in extant teleosts. GenBank/Ensembl
accession numbers for sequences in the MAFFT (Katoh and Standley, 2014) alignment: zebrafish: Prg4a,
NP_997918, Prg4b, XP_005160745; stickleback: Prg4a, ENSGACP00000019616, Prg4b, concatenate of
ENSGACP00000009961/ENSGACP00000009951/ENSGACP00000009946; medaka: Prg4a,
ENSORLP00000017289, Prg4b, ENSORLP00000011688; Spotted gar: Prg4, XP_015210531, Vtn,
XP_006641129; bowfin: Prg4, AAC_TPR.1.1 [http://phylofish.sigenae.org/]; human: PRG4, NP_005798,
VTN, NP_000629; mouse: Prg4, NP_067375; chicken: Prg4, ENSGALP00000008204; Anole lizard:
Prg4, XP_008112908; Western clawed frog: Prg4, XP_012817383; coelacanth: Prg4,
ENSLACP00000017445. (B) Orthologous pairwise synteny cluster from the Synteny Database (Catchen
et al., 2009) (window size: 50 genes) shows extensive conserved synteny of Prg4 gene regions on human
chromosome Hsa1 and spotted gar Loc10, supporting their orthology. (C) Composite cluster from the
Figure 4. 5. Evolution of vertebrate Proteoglycan 4 (Lubricin).
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Synteny Database (window size: 50 genes) compares the prg4 gene regions in zebrafish to the TGD
outgroup spotted gar. Double conserved synteny of prg4 on gar Loc10 and zebrafish prg4a on Dre2 and
prg4b on Dre20, supports the prg4 duplication as result of the TGD.
Requirement of zebrafish prg4b for adult maintenance of joints
A major function of the synovial cavity is to lubricate the joint, with loss of lubrication
resulting in age-related joint degeneration. We therefore asked whether expression of Prg4
orthologs by cells lining synovial-like joints in ray-finned fishes reflects a conserved requirement
of Lubricin protein in lubricating and hence maintaining these joints. To do so, we used TALE
nucleases (Huang et al., 2011; Sander et al., 2011) to generate loss-of-function deletion alleles
for zebrafish prg4a and prg4b (Figure 4.6A). Mice lacking Prg4 function (Koyama et al., 2014;
Rhee et al., 2005) and humans with homozygous loss of PRG4 in Camptodactyly-arthropathy-
coxa vara-pericarditis syndrome (Marcelino et al., 1999) have a progressive deterioration of joint
surfaces that includes loss of articular chondrocytes, accumulation of acellular matrix in the
cavity, synovial hyperplasia, and thickening of the deep chondrocyte layer. Consistent with the
lack of early joint defects in Prg4-/- mice, zebrafish doubly mutant for prg4a and prg4b had no
defects in the jaw joint at either 6 dpf or 1 mpf and no gross defects of the adult skeleton (Figure
4.6B-D). However, prg4a-/-; prg4b-/- zebrafish began to display a weak accumulation of
acellular matrix in limited domains of the joint surface at 2 mpf (Figure 4.6E), a phenotype that
became more severe by 6 mpf (Figure 4.6F). By 12 mpf, prg4a-/-; prg4b-/- zebrafish had
multiple jaw joint abnormalities, including an acellular matrix at the joint surface, synovial
hyperplasia, increased numbers of deep chondrocytes, and in some cases complete erosion of the
joint surface accompanied by underlying bone defects (Figure 4.6G, Figure 4.7, and additional
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examples in Figure 4.8). Quantification of jaw joint defects using an Osteoarthritis Research
Society International (OARSI) scoring system (Pritzker et al., 2006) that we modified for
zebrafish (Figure 4.8) confirmed that joint defects increased in severity during aging (Figure
4.6H), consistent with the progressive arthritis seen in Prg4-/- mice (Koyama et al., 2014; Rhee
et al., 2005) and CACP patients (Marcelino et al., 1999). Defects found in prg4a-/-; prg4b-/-
double mutants were also not confined to the jaw joint, appearing in the ray-scapula and inter-
radial joints of the pectoral fin (Figure 4.9A-C). Further, consistent with only prg4b being
enriched in jaw joint chondrocytes, prg4b but not prg4a single mutants displayed ray-scapula fin
joint defects and a similar severity of jaw joint defects as double mutants at 12 mpf (Figure
4.6G,H and Figure 4.9B). In contrast, we detected no changes in the hyoid joints of the face and
the intervertebral discs in 12 mpf prg4a-/-; prg4b-/- mutants (Figure 4.9D,E), consistent with
these joints lacking synovial cavities and high-level prg4b expression. Our data therefore support
a specific requirement for zebrafish Prg4 homologs in the adult maintenance of joints with
synovial-like morphology.
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Figure 4. 6. Progressive deterioration of the jaw joint in zebrafish lacking prg4b.
(A) Schematics of prg4a and prg4b TALEN mutants show deleted sequences. (B) X-ray imaging shows
no gross morphological bone defects of prg4a-/-; prg4b-/- mutants (DKO) at 12 mpf. (C) Alcian Blue
staining shows normal facial cartilages in DKO at 6 dpf. (D) SafraninO staining shows a normal jaw joint
between Meckel’s (M) and palatoquadrate (pq) cartilages in DKO at 1 mpf. (E-G) SafraninO staining at 2,
6, and 12 mpf shows increasingly abnormal joints between the jaw anguloarticular (aa) and quadrate (q)
bones in DKO. Defects include acellular matrix accumulation in the cavity (asterisks, see magnified
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regions in E’-G’) at all stages, and synovial hyperplasia (arrowheads) and expanded deep chondrocytes
(arrows) at 12 mpf. Single prg4b but not prg4a mutants showed similar jaw joint defects to DKO at 12
mpf. (H) Quantification of jaw joint defects using our modified OARSI system for zebrafish. Scale bar 50
μm, except in (B). See also Figure 4.7 and Figure 4.8.
96
SafraninO staining of a serial series of 5 μm sections were imaged every third section to capture a
complete wild-type and prg4a-/-; prg4b-/- (DKO) jaw joint at 12 mpf. See also Figure 4.6G. Scale bar,
50μm.
Figure 4. 7. Serial sections through a representative wild-type and prg4a-/-; prg4b-/- mutant jaw joint.
97
Figure 4. 8. OARSI scoring system for zebrafish.
SafraninO-stained histological sections demonstrate the defining features for each grade of joint damage
at the zebrafish jaw. Grade 0 – a smooth intact surface with normal superficial and deeper cartilage layers
(12 mpf, wild-type). Grade 1 – an uneven surface cartilage with small fibrillations limited to the
superficial layer (12 mpf, wild-type). Grade 2 – the superficial layer is disrupted with focal fibrillations
and some matrix loss (12 mpf, DKO). Grade 3 – vertical clefts or fissures extend beyond the superficial
layer, disrupting the deeper cartilage (12 mpf, DKO). Grade 4 – erosion of the superficial layer of
cartilage with matrix loss extending into deeper cartilage layers (12 mpf, DKO). Grade 5 – cartilage is
completely lost, exposing the bone surface (12 mpf, prg4b-/-). Grade 6 – the exposed bone surface is
deformed, showing altered contour of the joint (12 mpf, DKO). Individual grade and stage values were
determined for both the anterior and posterior sides of the left and right jaw joints from 3 histological
sections per joint. Overall joint OA scores were generated as an average of (grade) x (stage) for each
animal.
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Figure 4. 9. Requirement of prg4 gene function for fin joints of zebrafish.
(A) Schematic of pectoral fin joints. (B-E) SafraninO staining at 12 mpf shows abnormal joints between
the ray (r) and scapula (sc) joint of the pectoral fin (B) in prg4b but not prg4a single mutants and prg4a;
prg4b double mutants (DKO), and defects in the proximal radial (pr) and distal radial (dr) joint of the
pectoral fin (C) in DKO. Defects include acellular matrix accumulation in the cavity (asterisks), synovial
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hyperplasia (arrowheads), and expanded deep chondrocytes (arrow). In contrast, the hyoid joint (D) and
intervertebral discs (E) were normal in DKO. Phenotypes were consistently observed in three animals of
each genotype. Scale bar 50 μm.
4.4 Discussion
Although we do not know if Prg4b protein is secreted into joint cavities in zebrafish, or
whether the cavities are fully enclosed as in mammals, the finding that the jaw and fin joints of
zebrafish require the molecular lubricant Lubricin for their maintenance supports zebrafish
having synovial-like lubricated joints. Contrary to existing dogma (van der Kraan, 2013b), our
data suggest that lubricated synovial joints evolved before vertebrates moved onto land in at least
the common ancestor to all bony vertebrates. Although fish are not subject to gravity in the same
way as tetrapods, the evolution of synovial joints in fish-like ancestors may have facilitated
movement of first the jaw, and then the fins, against water resistance. Interestingly, recent studies
show that the radial joints of the pectoral fin, which we find to have synovial morphology and
prg4b dependency in zebrafish, are homologous to those of the tetrapod wrist (Gehrke et al.,
2015). The existence of these synovial joints at the base of the pectoral fins of ancestral bony fish
may therefore have facilitated their later functional evolution into the larger synovial joints of
tetrapod limbs. Our findings show that zebrafish can be a relevant model to understand the
development of synovial specializations, including the poorly understood process of cavitation.
The establishment of the first genetic model of arthritis in zebrafish will also allow a better
understanding of the developmental progression of synovial joint disease. In the future, it will be
exciting to test whether the articular cartilage lining synovial joints, which is affected in arthritis,
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displays the same regenerative potential as many of the other tissues in zebrafish (Knopf et al.,
2011; Morgan, 1901; Poss et al., 2002).
4.5 Materials and Methods
Fish husbandry and zebrafish lines
The Institutional Animal Care and Use Committees of the University of Southern
California, the University of California, Berkeley, and the University of Oregon approved all
zebrafish (Danio rerio), three-spined stickleback (Gasterosteus aculeatus) and spotted gar
(Lepisosteus oculatus) procedures, respectively – (IACUC #10885). Previously reported
zebrafish lines used in this study include Tg(sox10:dsRED)
el10
(Das and Crump, 2012),
trps1
j127aGt
(RRID:ZFIN_ZBD-GENO-100809-11) (Talbot et al., 2010), and
Tg(col2a1aBAC:GFP)
el483
(Askary et al., 2015).
Phylogenetic tree
The phylogenetic tree was generated using phyloT (http://phylot.biobyte.de/) with NCBI
taxonomy, and visualized in iTOL:Interactive Tree of Life (http://itol.embl.de/).
Histology
Adult zebrafish samples were fixed in 4% PFA at 4°C for 7 days. Following fixation,
animals were cut into smaller pieces to facilitate embedding and sectioning of the desired
structures. The samples were then decalcified in 20% EDTA solution for 10 days at room
temperature. For embedding, the tissue was first dehydrated through a series of ethanol washes
101
(30, 50, 70, 95, and 100%) for 20 min each. Then ethanol was replaced with Xylene substitute
Hemo-De (Electron Microscopy Sciences) in a series of 15 min washes (50, 75, and 100%
Hemo-De). Samples were then incubated in a 1:1 Hemo-De:paraffin (Paraplast X-tra, VWR)
solution at 65°C for an hour before an overnight incubation in 100% paraffin at 65°C. The
following day, samples were embedded in freshly melted paraffin. Larval and juvenile samples
were prepared following the same general procedure with shorter fixation and decalcification
steps (2 days fixation and 4 days decalcification). Animals younger than 3 weeks did not require
decalcification. Juvenile stickleback and spotted gar samples were processed as above for adult
zebrafish with the following changes: stickleback were decalcified in 20% EDTA for 7 days at
room temperature; isolated spotted gar heads were fixed in 4% PFA at 4°C for 10-14 days and
decalcified in 20% EDTA for 14 days at room temperature. For Hematoxylin & Eosin (H&E)
staining, 5 μm paraffin sections were deparaffinized with xylene and re-hydrated through an
ethanol series to distilled water. Sections were then stained in hematoxylin (VWR) for 2 min
followed by brief acetic acid rinse and 2 min in Blueing Reagent solution (VWR). 2 x 30 s
washes in water, 2 x 30 s in 95% ethanol. Sections were stained in Eosin (VWR) for 30 s
followed by 3 x 1 min washes in 95% ethanol and 2 x 1 min washes in 100% ethanol. Following
2 x 2 min in Hemo-De, samples were mounted with cytoseal (Richard-Allan Scientific) for
imaging. For Trichrome staining, 5 μm paraffin sections were deparaffinized with xylene and re-
hydrated through an ethanol series to distilled water. Trichrome stain was performed according
to manufacturer’s instructions using the Trichrome, Gomori One-Step, Aniline Blue Stain
(Newcomer Supply). For SafraninO staining, 5 μm paraffin sections were deparaffinized with
xylene and re-hydrated to distilled water. They were then stained in Weigert’s Iron Hematoxylin
(Newcomer Supply) for 5 min, washed in distilled water for 5 changes, differentiated in 0.06 N
HCl solution in 70% ethanol for 2 s followed by 3 more washes in water. Sections were then
102
stained in 0.02% Fast Green FCF (Sigma) for 1 min and rinsed for 30 s in 1% acetic acid.
Staining in 1% SafraninO (Newcomer Supply) was then performed for 30 min, followed by 3 x 1
min washes in 95% ethanol. Slides were then washed 2 x 1 min in 100% ethanol and 2 x 2 min in
Hemo-De, before mounting with cytoseal for imaging. For Alcian staining, juvenile zebrafish
were processed as described (Walker and Kimmel, 2007), and adult zebrafish were processed
using a modified version of the protocol
(https://wiki.zfin.org/pages/viewpage.action?pageId=13107375).
In situ hybridization
Probes were generated for zebrafish prg4a, prg4b, has3, acana, col10a1, and matn1 using
PCR amplification from cDNA. See Supplemental file 1A for primers used in amplification of
zebrafish probe templates. Probes for stickleback prg4a and prg4b and gar prg4 were
synthesized from gBlocks Gene Fragments (IDT). The DNA sequences used for probe
generation are listed in Supplemental file 1B. Stickleback prg4b sequence was determined
through sequence homology comparison with zebrafish Prg4b protein sequence using BlastP
search in Ensembl. Two independent probes were designed for the putative stickleback prg4b
(ENSGACG00000007505; ENSGACG00000007501) and both showed the same expression
pattern in three independent animals. gBlocks or PCR products from cDNA amplification were
subcloned into pCR-Blunt II TOPO vector (Invitrogen). Following sequence confirmation,
digoxigenin (DIG)-labelled antisense probes were synthesized using T7 or SP6 RNA polymerase
(Roche). In situ hybridization protocol was modified from (Lien et al., 2006). In brief, after de-
paraffinization, slides were digested in 7.5 µg/ml proteinase K for 5 min and fixed in 4%
PFA/0.2% gluteraldehyde for 20 min. Each slide was incubated overnight at 65˚C with 1 ug of
DIG-labelled riboprobe diluted in hybridization buffer. After hybridization, slides were washed
103
three times in 1x SSC/Formamide at 65˚C for 30 min, and three times in MABT for 15 min.
Following 1 hr of blocking in 2% Blocking Buffer (Roche), hybridization was detected with anti-
DIG-AP antibody (Roche, RRID:AB_514497)) and developed with NBT/BCIP substrate
colorimetric reaction (Roche). Slides were counterstained with nuclear fast red (Vector
Laboratories) prior to mounting.
Immunofluorescence
Paraffin sections (5 µm) were de-paraffinized with xylene and rehydrated through an
ethanol series to 1xPBST. Antigen retrieval was performed using pH 6.0 sodium citrate buffer in
a steamer for 35 min. Sections were blocked with 2% donkey serum in PBST for 30 min at room
temperature. Sections were incubated with primary antibodies against Col2a1 (goat polyclonal,
SC7763, Santa Cruz, RRID:AB_2229686) and Aggrecan (Cat# 13880-1-AP, Proteintech)
overnight at 4˚C. Sections were further incubated in secondary antibodies and Hoechst 33342
nuclear stain for 1 hour at room temperature prior to mounting with Fluoromount-G (Southern
Biotech).
Imaging
Fluorescence imaging of live animals and sections was performed using a Zeiss LSM5
confocal microscope. Histology and colorimetric in situ hybridization slides were photographed
using a Leica D8 2500 microscope. Alcian Blue-stained samples were imaged using a Leica
S8APO microscope. For microCT (µCT), adult zebrafish were euthanized and fixed in 4% PFA
overnight and rinsed twice in 1x PBS. The fish head was dissected and glued to a Pasteur pipette
to place it next to the scan head. The scans were performed in air on a XT H 225S T µCT
scanner (Nikon Metrology, Brighton, MI) with a PerkinElmer 1621 detector at 120 kVp, 26 uA
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500 ms exposure time resulting in an isotropic 3 micron voxel volume. A molybdenum target
was used with no additional filtration of the beam. Raw data was reconstructed in CT Pro 3D
v4.3.4 (XT Software Suite, Nikon Metrology) and video rendering performed on VG StudioMax
v2.2 (Volume Graphics GmbH, Heidelberg, Germany). X-ray imaging of fixed adult zebrafish
utilized an UltraFocus
60 x-ray cabinet (Faxitron Bioptics, Tucson, AZ).
Construction of Prg4 knockout alleles
To make knockout alleles for prg4a and prg4b, two pairs of Transcription Activator-Like
Effector Nucleases (TALENs) were used per gene to remove most of the coding sequence of the
target genes. TALENs were designed and constructed as described previously (Sanjana et al.,
2012). TALEN pairs were targeted toward the following sequences in the zebrafish genome:
TTGGTCTCTTCTGGCTCTGC and TCGTCTGCTGCTCAGGGTGA for prg4a 5’ TALENs,
TAGGCGTCCCGTCACCCATT and CGCTGCAACTGCCAGGGCAA for prg4a 3’ TALENs,
TGCTGTTTGTGTGGGTCTCC and CACGTAAGCCAACAGATCGA for prg4b 5’ TALENs,
and TCCCCCAGCTGCAGCACTGG and TCTCACGAACCTGGAGAGGA for prg4b 3’
TALENs. ARCA-capped RNA for each TALEN was transcribed in vitro using the mMESSAGE
mMACHINE® T7 Ultra Kit (Life Technologies). The RNA for four TALENs corresponding to
each gene were mixed and injected into one-cell stage embryos to generate founder lines.
Founders were then outcrossed to wild-type fish and the progenies were tested for genomic
deletions. Deletion alleles were identified in the F1 generation by cloning and sequencing of
amplicons that span the TALEN target sites for each gene. We obtained single prg4a
el687
(7686
bp deletion) and prg4b
el594
(11,638 bp deletion) alleles by screening the progeny of 6 and 21
injected animals, respectively. For the list of genotyping primers see Supplemental file 1C.
105
Zebrafish OARSI scoring system
5 µm serial sections throughout the jaw joint were stained with SafraninO and imaged as
described above. For both the left and right jaw joint, three representative images were selected
for each sample. Care was taken to select images that represent equivalent sectioning planes in
wild-type and mutant joints. Grade (0-6) and stage (0-4) values were assigned to the anterior
(anguloarticular) and posterior (quadrate) surfaces in each of the six joint images for each
sample. The grade and stage value for each individual articular surface were multiplied to
generate the score for that joint surface. Scores for both surfaces of the left and right joints were
then averaged for each animal.
Statistical analysis
Data are presented as a scatter plot with a line for the mean. Statistical analysis of average
OARSI scores was performed using GraphPad Prism 7 (RRID:SCR_002798), with a two-tailed
Student’s t-test used to generate the P values.
4.6 Credits
This study has been the result of a collaborative and shared effort between the author and Joanna
Smeeton in Gage Crump lab, who is contributed equally to this work. Other co-authors include
Sandeep Paul, Simone Schindler, Ingo Braasch, Nicholas A. Ellis, John Postlethwait, Craig T.
Miller, and Gage Crump.
106
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Koyama, E., Saunders, C., Salhab, I., Decker, R.S., Chen, I., Um, H., Pacifici, M., and Nah, H.D. (2014). Lubricin is
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108
Conc l us i on
Joints provide flexibility to a rigid skeleton. For a developmental biologist, joints are the
product of precise coordination of many signaling pathways, most notably Bmp, Wnt, and Hh,
that together specify the position of the joint within the skeletal condensation, guide the cells in
that position to make very different fate decisions compared to their neighboring skeletal
precursors, and seamlessly integrate the resulting cell types into a functional organ. This
complicated process is also surprisingly robust and reliable; consequently, congenital anomalies
of joints are uncommon. On the other hand, long term maintenance of the load bearing joints
often fails, as a result osteoarthritis, the progressive degeneration of joints, is a major health
concern that affects approximately 27 million Americans. The clinical situation provides a strong
motivation to study the development of the cell types of the joint, especially the articular
cartilage, and potential mechanisms for joint regeneration.
Joints are also interesting from an evolutionary point of view. The joints of the
craniofacial skeleton enabled the ancestors of jawed vertebrates to use muscle force for active
and efficient opening and closing of the mouth, which resulted in much improved ventilatory
function and predatory ability. Whereas in the jawless ancestors of vertebrates, an inspiratory
phase of ventilation was likely achieved by passive recoil of unjointed branchial arches, as in
jawless vertebrates today, jawed vertebrates could use the muscles attached to their jointed
highly-mobile jaws to open the mouth during forceful inspiration and close it to prevent leakage
in a forceful expiration. At the same time, this enabled the jawed vertebrates to catch evasive
prey, sucking the prey into the mouth during a forceful inspiration and then clamping onto the
prey during expiration (Mallatt, 2008; Mallatt, 1996). Despite the fact that joints were
instrumental in evolutionary success of jawed vertebrates, we know little about how and when
109
different types of joints evolved, and how they acquired the sophisticated specializations we see
today.
Here, we demonstrated that a study of zebrafish joints can help further our understanding
of the development and evolution of joint. The zebrafish hyoid joint consists of chondrocytes
that initiate a cartilage differentiation pathway, turning on the expression of SoxE transcription
factors, but subsequently block overt differentiation. Preventing overt cartilage differentiation
and hypertrophy is also a major step in regulation of cell fate in the interzone of mammalian
synovial joints. Therefore, the hyoid joint of the fish shares critical features with considerably
more complex mammalian joints.
In contrast to a complex mammalian joint such as the knee, the zebrafish hyoid joint is
composed of only one cell type. Therefore, it provides a simpler and more tractable model to
study factors involved in inhibition of cartilage differentiation in joints. We utilized this to find
out the role of Iroquois transcription factors in joint development. Although the expression of
Iroquois family members, Irx1 and Irx2, in mouse digit joints was known for more than a decade
(Zulch et al., 2001), their function had remained elusive, partly due to the functional redundancy
of Irx genes. We showed that irx genes in zebrafish, specifically irx7 and irx5a, halt cartilage
differentiation program in joint cells through direct repression of cartilage differentiation genes,
such as col2a1a.
This function is conserved in the mouse chondrogenic cell line, ATDC5. In this line,
ectopic expression of either mouse Irx1, or zebrafish irx7, inhibits cartilage differentiation. A
promising future research direction will be to understand the extent to which the pathways that
establish joint cartilage fate during development contribute to maintaining this fate over the life-
span of the animal. In the case of zebrafish hyoid joint, irx7 expression is maintained beyond the
110
embryonic and early larval stages. However, due to the early joint phenotype in irx7 mutants, it
is unclear whether there is continuous requirement for irx7 to inhibit cartilage differentiation at
the joint surface.
Future studies will aim to explore the function of novel joint specific candidate genes,
identified through unbiased expression analysis of the pharyngeal arches. One important
unanswered question is how signaling through members of the Tgfβ superfamily (i.e. Bmp, Gdf,
and Tgfβ) is fine-tuned in the joints to specify individual joint cell types, given that some family
members promote differentiation of transient cartilage and stimulate bone formation in
neighboring skeletal elements. Through our genomic study, we discovered that fsta, an inhibitor
of Tgfβ signaling not previously implicated in joint development, is expressed in the joint
forming domain of the pharyngeal arches. Further analysis of fsta function in this context can
help us better appreciate the complexity of Tgfβ signaling regulation in joint development.
Concerning the evolution of the joints, we provided substantial histological and
molecular evidence that certain joints (e.g. jaw and pectoral fin radial joints) in distant ray finned
fish species acquire definitive characteristics of tetrapod synovial joints during late larval and
adult stages. This shows that lubricated synovial joints evolved much earlier than previously
thought, and were likely present in the ancestors of all bony vertebrates. We were not able to
extend this study beyond bony vertebrates, to cartilaginous fish, due to a paucity of samples and
genomic data. So whether synovial joints are in fact present in cartilaginous fish remains to
addressed in future. If so, the origins of synovial joints may be closely linked to the evolution of
jaw itself.
Beyond the evolutionary implications, the discovery of lubricated synovial joints in
zebrafish is significant from the perspective of developmental biology and regenerative
111
medicine. In chapter 4, we showed two examples of how zebrafish can be useful as a synovial
joint model: observing cavitation of the joint in live animals and modeling certain human
synovial joint conditions, namely CACP syndrome. As the zebrafish also has the ability to
regenerate many tissues and organs, including fin (Gemberling et al., 2013), heart (Poss et al.,
2002), and jaw bone (Paul et al., 2016), an obvious next step is to determine if the zebrafish can
regenerate damaged synovial joints. In this case, the zebrafish will offer a unique opportunity to
study joint regeneration.
References
Gemberling, M., Bailey, T.J., Hyde, D.R., and Poss, K.D. (2013). The zebrafish as a model for complex tissue
regeneration. Trends in genetics : TIG 29, 611-620.
Mallatt, J. (2008). The origin of the vertebrate jaw: neoclassical ideas versus newer, development-based ideas.
Zoological science 25, 990-998.
Mallatt, J.O.N. (1996). Ventilation and the origin of jawed vertebrates: a new mouth. Zoological Journal of the
Linnean Society 117, 329-404.
Paul, S., Schindler, S., Giovannone, D., de Millo Terrazzani, A., Mariani, F.V., and Crump, J.G. (2016). Ihha
induces hybrid cartilage-bone cells during zebrafish jawbone regeneration. Development 143, 2066-2076.
Poss, K.D., Wilson, L.G., and Keating, M.T. (2002). Heart regeneration in zebrafish. Science 298, 2188-2190.
Zulch, A., Becker, M.B., and Gruss, P. (2001). Expression pattern of Irx1 and Irx2 during mouse digit development.
Mech Dev 106, 159-162.
Abstract (if available)
Abstract
We owe the flexibility of our bodies to the joints that connect our bones to each other. Proper development of these joints requires the integration of multiple tissue types, made possible by coordination of several signaling and regulatory pathways. Conceptually, the process of joint development can be divided into three steps: positioning of the joint within the skeleton, specification of joint cell identities, and formation of joint specializations. Positioning the prospective joint region involves establishment of an “interzone” region of joint progenitor cells within a nascent cartilage condensation. Inside the interzone, tight regulation of several signaling pathways, most notably Bmp and Tgfβ, guides the joint chondrocytes through their cell fate decisions and prevents their hypertrophic maturation. Joint cells then acquire further specializations according to the type and function of each joint. ❧ The majority of research on joint development, so far, has been conducted using mice and chicks as model organisms. We believe that zebrafish can offer a complementary reductionist approach towards understanding joint development. As an example, we used the hyoid joint of zebrafish, which functions in gill ventilation, to gain new insights into the regulation of cell identity in joints. At the hyoid joint, prospective joint chondrocytes express SoxE transcription factors, sox9 and sox10. However, in contrast to the differentiating chondrocytes in cartilage condensations, joint chondrocytes make only low levels of Col2a1a and Aggrecan. We found that members of Iroquois transcription factors, irx7 and irx5a, are expressed in the hyoid joint chondrocytes. We then generated irx7 and irx5a null mutants and showed that mutant hyoid joint chondrocytes inappropriately mature into high Col2a1a-expressing chondrocytes, in the absence of direct repression of a col2a1a enhancer by Iroquois proteins. In mammals, Irx1 and Irx2 are expressed in the developing interphalangeal joints. We showed that either mouse Irx1 or zebrafish Irx7 can repress the chondrogenic differentiation of a murine chondrogenic cell line. This finding suggests that the function of Irx genes in inhibiting excessive differentiation of joint cells may be conserved from zebrafish to mouse. ❧ Additionally, we took an unbiased genomic approach to find more regulators of cell fate specification and patterning within zebrafish facial skeleton. The Facial skeleton is formed by neural crest derived cells of the pharyngeal arches. Prior to condensation of arch mesenchymal cells, they are patterned into at least three domains along the Dorsal-Ventral (DV) axis. Whereas dorsal and ventral domains of the arches give rise to dorsal and ventral elements of the facial skeleton, joints are formed by cells from the intermediate domain. We used transgenic zebrafish lines that label different domains, or combinations of domains, within pharyngeal arches to sort and sequence cells, to identify their transcriptome. We then used our expression data to search for genes that are enriched in each domain. Our assumption is that the genes which are important for specification of each domain must be expressed preferentially in those domains. This analysis has provided us with novel candidates, for in depth genetic and functional analysis. ❧ At birth, major joint defects are relatively uncommon. However, the progressive degeneration of joints in osteoarthritis is the leading cause of disability in the United States. Consequently, among all the different types of joints, the study of synovial joints, the type that is prone to osteoarthritis, is arguably of highest clinical relevance. Synovial joints, such as those in our limbs, are the most sophisticated and most flexible types of joints. They are composed of multiple tissue types: permanent cartilage that cushions the articulating bones, synovial membranes that enclose a lubricating fluid-filled cavity, and a fibrous capsule and ligaments that provide structural support. It is commonly believed that synovial joints evolved when vertebrates transitioned from water to land, because lubrication could facilitate movement of their load bearing joints which were exposed to increased gravitational pressure on land. Consistently, at the stage that they are best studied, i.e. early larval stages, none of the zebrafish joints resemble the lubricated synovial joints of mammals closely. However, we provide substantial evidence that certain joints in zebrafish body, including the jaw joint, start to acquire features of synovial joints at later stages. Building on histological observations that the jaw joints of diverse fish species (e.g. spotted gar, stickleback, and zebrafish) have cavities, synovial membranes, articular cartilage, and fibrous capsules, we showed that these joints also produce Prg4/Lubricin, a lubricating protein which also serves as a marker of synovial joints. Further, we generated mutant zebrafish lacking the Lubricin-encoding gene, prg4b, and showed that they develop arthritic changes to their jaw joint that closely resemble those observed in mutant mice and human patients lacking Lubricin. Synovial properties in zebrafish are not limited to the jaw joint, as joints at the base of the pectoral fin also produce and require Lubricin for their maintenance. These findings show that synovial joints were present in the common ancestor of all bony vertebrates. Particularly, the Lubricin mutant establishes the first genetic arthritis model in zebrafish, enabling the study of synovial joints in this powerful and highly regenerative model. Future work in zebrafish has the potential to address unanswered questions such as the cellular mechanism of cavitation and the ability of synovial joints to regenerate.
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Askary, Amjad
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Core Title
Development and evolution of skeletal joints: lessons learned from studying zebrafish
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Keck School of Medicine
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Doctor of Philosophy
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Genetic, Molecular and Cellular Biology
Publication Date
09/23/2016
Defense Date
07/19/2016
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