Fish - Semantic Scholar

DDMOD-408; No of Pages 9

Drug Discovery Today: Disease Models

DRUG DISCOVERY

TODAY

DISEASE

MODELS

Vol. xxx, No. xx 2014

Editors-in-Chief Jan Tornell – AstraZeneca, Sweden Andrew McCulloch – University of California, SanDiego, USA

Bone biology in animal models, including testing of bone biomaterials

Fish: a suitable system to model human bone disorders and discover drugs with osteogenic or osteotoxic activities Vincent Laize´1,3, Paulo J. Gavaia1,3, M. Leonor Cancela1,2,3,* 1 2

Centre of Marine Sciences, University of Algarve, 8005-139 Faro, Portugal Department of Biomedical Sciences and Medicine, University of Algarve, 8005-139 Faro, Portugal

This review discusses the suitability and advantages of teleost fish for studying underlying mechanisms of normal and pathological development and mineralization of vertebrate skeleton, presents a selection of zebrafish mutants and transgenic lines modeling hu-

Section editors: Oskar Hoffmann – Pharmacology and Toxicology, UZA II Pharmacy Center, Vienna, Austria. Wilhelm Hofstetter – Clinical Research Department, University of Bern, Bern, Switzerland.

man skeletal diseases and highlights currently available fish systems for identifying and characterizing novel osteogenic and osteotoxic molecules. Introduction The last decade has seen an exponential interest in the use of model organisms capable of providing suitable alternatives to mammalian models and allowing quicker and less expensive approaches, leading to significant improvements in our knowledge on the molecular basis of human pathologies. In this respect, the study of skeletal diseases has not been an exception but requires the use of vertebrates for obvious reasons. Fish, in particular zebrafish and medaka, have been used with a growing success due to their many similarities with mammals in molecular pathways and mechanisms involved in the onset of patterning and development of skeletal structures. Many reports have confirmed the suitability of fish systems to model many of the pathological defects affecting human skeletal formation, osteoclastic bone resorption, osteoblastic bone mineralization and extracellular matrix maintenance, but also to visualize the onset of normal and *Corresponding author.: M.L. Cancela ([email protected], [email protected]) 3 All authors contributed equally. 1740-6757/$ ß 2014 Elsevier Ltd. All rights reserved.

abnormal skeletal structures, and the possibility of rapidly detecting drug-induced osteogenic or osteotoxic effects in phenotype-based assays.

Teleost fish and human skeletons are remarkably similar Despite some small differences that can be attributed to evolutionary distance (their last common ancestor existed approximately 420 million years ago), anatomic and developmental features of teleost fish and mammalian skeletons are remarkably similar, with much of the skull, axial and appendicular skeleton formed by identical bones, and with a high conservation of the developmental events and underlying mechanisms of skeletogenesis, including the early formation of a cartilaginous anlage followed by bone formation through endochondral and dermal ossification [1,2]. Main differences and similarities between fish and mammalian bone are summarized in Table 1 (see [3] for a more comprehensive comparison). The most striking difference is probably the occurrence of acellular bone (devoid of osteocytes) and mononucleated osteoclasts in most teleost fish species, while mammals have exclusively cellular bone and multinucleated osteoclasts. Interestingly, osteocytic bone and multinucleated osteoclast

http://dx.doi.org/10.1016/j.ddmod.2014.08.001

e1

Please cite this article in press as: Laize´ V, et al. Fish: a suitable system to model human bone disorders and discover drugs with osteogenic or osteotoxic activities, Drug Discov

DDMOD-408; No of Pages 9 Drug Discovery Today: Disease Models | Bone biology in animal models, including testing of bone biomaterials


Table 1. Features of fish and human bone: similarities and differences Fish skeleton

Human skeleton

Endochondral and membranous ossification Hydroxyapatite-like calcium-phosphate crystals Bone formation by osteoblasts positive for alkaline phosphatase Bone resorption by osteoclasts positive for tartrate-resistance acid phosphatase Mesenchymal origin of osteoblasts; hematopoietic origin of osteoclasts Osteocytic and anosteocytic bone Osteocytic bone Dendritic processes in a well-developed lacunocanalicular system Dendritic processes mostly absent and no lacunocanalicular system Mono- and multinucleated osteoclasts Multinucleated osteoclasts Mechanosensing by osteoblasts and lining cells Mechanosensing by osteocytes Collagen I and II in bone Collagen I in bone Perichordal ossification of the vertebral column Endochondral ossification of the vertebral column 16 types of cartilage 3 types of cartilage

occur in zebrafish [4,5]. Also noteworthy is the fact that most teleost fish ossify the vertebrae directly over the notochord, while mammals have a cartilaginous precursor [6,7]. Fish also have a much higher diversity of cartilage types and intermediate tissues than what is found in adult mammals [4,8,9]. Despite those differences, the availability of fish mutants exhibiting features resembling human pathologies has contributed to validate fish as a model organism to study mechanisms underlying skeleton development and skeletal disorders. Among teleost fishes, zebrafish Danio rerio (Hamilton, 1822) has been the target of a growing interest from the scientific community. Zebrafish is a small freshwater fish from tropical regions of South and Southeast Asia for which numerous models of human diseases have been identified or developed during the last decade, leading to its recognition as a suitable model for biomedical research (reviewed in [10]). Zebrafish embryos develop externally and are transparent, allowing for direct observation of embryonic development and organ growth. Additional features such as small size (3– 4 cm long), high fecundity (mature females can lay hundreds of non-adhesive eggs every week), short generation time (adulthood attained in 3–4 months) and rapid development (most body structures are visible 48 h after fertilization), robustness (they are easy to manipulate, adapt to a wide range of environments and can be kept in large shoals) and availability of genetic techniques, have reinforced the attractiveness of the zebrafish as a laboratory model. Finally and importantly, the zebrafish genome has been almost entirely sequenced and annotated, with most human genes having orthologs in zebrafish [11]. In addition, it is estimated that approximately 70% of human disease genes have functional homologs in zebrafish [12]. The key regulators of bone formation have been highly conserved, and corresponding zebrafish orthologs share significant sequence similarities and overlapping of expression patterns [13]. For phenotype-based assays related to skeletogenesis, the transparency and external/rapid development of fish embryos is a clear e2

References [1,59] [1] [5] [5] [59] [5] [60–62] [63] [59,61,64] [1,8] [6] [8]

advantage. It is possible to monitor each stage of skeleton development and determine at high resolution the role of single cells in bone and cartilage formation/mineralization. However, zebrafish is not a mammalian species and the lack of some organs (e.g. lung or mammary gland) represents a limitation to the modeling of some human diseases. In addition, phenotypic characteristics of diseases caused by orthologous genes can differ between fish and human. Because it suffered the teleost-specific whole genome duplication event, zebrafish genome contains numerous paralogous genes that resulted in gene sub-functionalization and/or neofunctionalization, a situation that may limit, in some cases, the use of zebrafish as a disease model. However, this is also true in studies using the mouse model [11,14]. Still, zebrafish is a relevant model organism to study vertebrate development and human diseases, and is rapidly acquiring standard prominence worldwide as an alternative and complement to rodent models, which remain the standard system in pre-human drug tests.

Fish models of human skeletal disorders In the last decade, there has been an increasing effort in producing mutant and transgenic fish lines with the objective of modeling human skeletal disorders and improving the visualization of the skeleton. These achievements have decisively contribute to facilitate studies towards uncovering novel drugs with the potential to treat the many skeletal pathologies that affect millions of people worldwide and which incidence is growing due to the increase in human life span. Human diseases such as osteogenesis imperfecta, bone loss (osteoporosis and osteopenia), craniosynostosis, craniofacial defects and spinal deformities (scoliosis, holospondyly) are examples, among many others, of pathologies for which fish mutants are available and being used to unveil the corresponding changes in normal molecular pathways, thus contributing to discover therapeutic molecules capable of rescuing these pathologies (Table 2). Although medaka and guppy have also been used to model human bone disorders,

www.drugdiscoverytoday.com


DDMOD-408; No of Pages 9 Vol. xxx, No. xx 2014

Drug Discovery Today: Disease Models | Bone biology in animal models, including testing of bone biomaterials

Table 2. Example of fish models of human bone and skeletal disorders (see also [90]) Human bone/skeletal disorders (bone/skeletal phenotype)

Fish model systems

Affected gene(s)

References

Osteogenesis imperfecta (OI) (reduced bone density, bone fragility, skeletal deformities) Osteoporosis (reduced bone mineral density) Glucocorticoid-induced osteoporosis (GIOP) (reduced bone mineral density upon use of steroids) Iron-induced osteoporosis (reduced bone mineral density upon iron overload) Raine syndrome (RNS) (increased ossification) Multiple osteochondromas (MO) (cartilaginous bone tumors leading to skeletal deformities)

Zebrafish chihuahua (chi) mutant Zebrafish frilly fins (frf) mutant rankl-induced medaka

col1a1 bmp1 rankl

[65,66]

Mucolipidosis II (ML-II) (skeletal, craniofacial and joint abnormalities) Craniosynostosis (premature fusion of cranial sutures) Holospondyly (fusion of vertebral centra) Fibrodysplasia ossificans progressiva (FOP) (heterotopic endochondral ossification) Idiopathic scoliosis (spinal deformity) Arterial calcification of infancy (ectopic mineralization) Holoprosencephaly (HPE) (craniofacial defects) Campomelic dysplasia (craniofacial defects) Ehlers-Danlos syndrome (EDS) (craniofacial defects) DiGeorge syndrome (DGS) (craniofacial defects) Cranio-lenticulo-sutural dysplasia (CLSD) (craniofacial defects and short stature) Osteopathy related to mineral homeostasis (failure to form mineralized bone) Osteopathy related to abnormal ECM deposition (craniofacial defects) Osteopathy related to delayed mineralization (delayed vertebrae calcification) Hyperossification (hyperossification and skeletal overgrowth)

[67]

Prednisolone-treated zebrafish

[56,68]

Iron-overloaded zebrafish

[28]

Zebrafish fam2b mutant

fam20b

[69]

ext2 extl3 papst1 gnptab

[70,71]

[72]

Zebrafish dolphin (dol) and stocksteif (sst) mutants

cyp26b1

[73]

Zebrafish stocksteif (sst) mutant Retinoic acid-treated zebrafish Zebrafish lost-a-fin (laf) mutant

cyp26b1

[74]

acvr1/alk8

[75]

Guppy curveback mutant

Not determined

[76]

Zebrafish dragonfin (dgf) mutant

enpp1

[16]

Zebrafish sonic-you (syu) mutant

shh

[77]

Zebrafish jellyfish (jef) mutant

sox9a

[78]

Zebrafish b4galt7 mutant

b4galt7

[79]

Zebrafish van-gogh (vgo) mutant

tbx1

[80]

Zebrafish crusher (cru) mutant

sec23a

[81]

Zebrafish no bone (nob) mutant

entpd5

[16]

Zebrafish Zebrafish Zebrafish Zebrafish

creb3l2 uxs1 sec24d Not determined

[82–84]

[85]

rpz

[86]

Zebrafish Zebrafish Zebrafish Zebrafish

dackel (dak) mutant boxer (box) mutant pinscher (pic) mutant gnptab morphant

feelgood (fel) mutant man o’war (mow) mutant bulldog (bul) mutant bone calcification slow (bcs) mutant

Zebrafish rapunzel (rpz) mutant

most diseases models have been developed from zebrafish mutants. Among the first mutants developed and characterized, the zebrafish chihuahua, which lacks a functional type 1 collagen gene, exhibits a skeletal dysplasia resembling human osteogenis imperfecta characterized by extreme bone fragility. Zebrafish mutants sonic you – lacking a functional sonic hedgehog gene, which is a key regulator of vertebrate organogenesis – and jellyfish – lacking a functional sox9a gene, which is a transcription factor involved in chondrocyte differentiation – exhibit skeletal dysplasia resembling human craniofacial syndromes. Guppy is also a model and its mutant

curveback exhibits a skeletal dysplasia resembling idiopatic scoliosis. Following these first mutants, many more have been developed and characterized (Table 2) and its numbers are continuously growing. A simple search in bibliography using as key words ‘zebrafish’ and ‘human diseases’ can undoubtedly show the exponential interest in zebrafish in the last decade. Recently, zebrafish has also been proposed as a model of excellence to study the molecular and cellular events underlying the development of ectopic mineralizations, which are observed in a number of human pathologies such as vascular calcification, skin calcifications, renal www.drugdiscoverytoday.com

e3



calcium stones, among others [15]. For example, the dragonfish mutant shows a broad range of ectopic calcifications around perichondral bone in the craniofacial skeleton as well as in the brain, the neural tube and the heart. On the contrary, the no bone mutant fails to produce mineralized bone [16]. Furthermore, the growing number of transgenic lines expressing fluorescent proteins under the control of promoters of bone-related genes such as osx (sp7), oc2, runx2, sox9, sox10, barx1, col2 and col10 among others [17–21], provide a plethora of in vivo tools which allow studies with a level of morphological detail and at a temporal scale never achieved before. Thus it can be anticipated that these improvements, both technical and in availability of in vivo models with improved capabilities will lead to new discoveries at a much higher pace than before. Information on the availability of mutant and transgenic zebrafish lines can be found at the European Zebrafish Resource Center (EZRC; http:// www.ezrc.kit.edu) or at the Zebrafish International Resource Center (ZIRC; http://zebrafish.org).

Fish systems to discover osteogenic drugs Given the important advances in technology, public in general as well as public and private decision makers expect that these should accelerate the discovery of novel treatments, pressing those involved in biomedical research as well as pharma and biotech companies, to bring new drugs into the market. Thus, economical as well as political pressures require more and more the use of high throughput methods to achieve the expected results. Given the advantages highlighted above, zebrafish and medaka stand up among other fish as models of choice to be used in biomedical research, in particular for this quest for novel drugs suitable to be used in bone therapeutics. Its importance can be also measured by the growing number of articles focusing on this issue in the last decade [10,22–26]. Here we describe available systems used in screenings for osteogenic and mineralogenic drugs. The success of these approaches confirms that fish, in particular the cellular boned zebrafish, is an excellent model for discovering therapeutic molecules for human skeletal disorders (Fig. 1). Accordingly, Table 3 presents a number of relevant studies which used zebrafish as an in vivo model to uncover, following screening of various panels of drugs, novel osteogenic and/or mineralogenic drugs, or drugs capable of reverting pathological conditions, thus proving once more the importance of this model organism for discovering therapeutic molecules of interest for human skeletal disorders. For example, the recent screening of 320 compounds in the zebrafish mutant Snca1 led to the identification of clemizole as a potential treatment for Dravet syndrome [27]; in another study using zebrafish, deferoxamine was found to effectively counteract iron-overload-induced inhibition of osteogenesis [28]. Similarly, zebrafish has also been used as e4


a choice model to unveil the effects of pollutants and other toxic molecules on skeletogenesis, with high relevance for environmental monitoring and ecotoxicology.

Skeletogenesis in fish larvae Looking at the effect of a molecule in the whole-organism is a preferable approach since it not only allows the identification of potential pitfalls but also the determination of therapeutic activity, range of action and general toxicity. Fish, in particular zebrafish, embryos/larvae are small, transparent and available in large quantities, and thus can be easily accommodated in 96-well plates for high-throughput screening of molecule libraries. In zebrafish, the onset of bone formation and mineralization occurs as early as 2 day post-fertilization (dpf) and is first evident at 3 dpf in the cleithrum then in the branchial and cranial skeletons [29]. Furthermore, it can be assessed easily through whole-mount bone-specific staining due to the transparency of the larvae at these early stages. The fast development of the fish skeleton allows for short duration assays (few days), something difficult in traditional mammalian models. Larvae are initially grown in petri dishes then arrayed in 96-well plates at a density up to five fish per well. Molecules are added directly to the liquid medium in which the embryos/larvae develop. In wild-type strains, ossification is detected through alizarin red S staining in 6–11 dpf zebrafish larvae [30] and quantified from the area and intensity of staining determined from color analysis of ventral pictures of the head skeleton [31] (Fig. 1b). Zebrafish transgenic lines using fluorescent reporters highlighting particular structures in the skeleton can also be used to observe specific bone elements/cells and quantify their area/number without the need for euthanasia. For example, Tg(oc2:GFP; osx:mCherry) transgenic zebrafish line (Fig. 1c), where GFP and mCherry production is under the control of osteocalcin 2 and osterix promoters, respectively, can be used to determine the size of the operculum or the number of pre- versus mature osteoblasts. Similarly, col10a1:nlGFP transgenic zebrafish line, where GFP production is under the control of collagen 10 promoter, a marker of early osteoblastic precursors, can be used to assess events preceding mineralization in cranial and axial skeleton [21]. In addition to its common use as a colorant in bone research, Alizarin red S is also used as a fluorochrome label of mineralized tissues, often as a complement to fluorescent reporters in transgenic fish ([32]; Bensimon-Brito et al., in preparation). In all cases, osteogenic/ osteotoxic effects are determined from the comparative analysis of treated versus control fish.

Osteogenesis in regenerating teleost fish fin The caudal fin of teleost fish is remarkably simple, accessible and can be restored upon amputation or damage. Because of these characteristics, it has logically become an excellent system for investigating underlying mechanisms of epimorphic





(a)

Skeletal deformities Regenerating fin rays

Developing operculum

Mineralizing scales

Mineralogenic vertebraderived cell lines

11 dpf

(b)

Cl

Tg(oc2:GFP; osx:mCherry) 16 dpf

(c)

Op Operculum Nc PT

Ps Cb5

(d)

(e)

Lepido -trichia

(f)

AR-S stained nodules

5 dpa Drug Discovery Today: Disease Models

Figure 1. (a) Fish systems used to study/screen molecules for osteogenic effects. (b) Ventral view of whole-mount alizarin red S (AR-S) – alcian blue (AB) stained zebrafish larvae at 11 days post-fertilization (dpf). Calcified structures appear in red; area and intensity of staining is determined for the notochord (Nc), operculum (Op), parasphenoid (Ps) cleithrum (Cl), ceratobranchial 5 (Cb5) and pharyngeal teeth (PT). (c) Lateral view of Tg(oc2:GFP; osx:mCherry) transgenic zebrafish at 16 dpf. Dotted white line indicates operculum area. (d) Lateral view of AR-S stained regenerating caudal fin of a juvenile zebrafish. Black triangle indicates the amputation plan; Dotted black line indicates regenerated area; Solid black line indicates area of new bone formation. (e) AR-S-stained elasmoid scale from the dorsal region of a juvenile gilthead seabream (Sparus aurata L.). (f) AR-S stained mineral nodules deposited within the extracellular matrix of gilthead seabream VSa16 cell line (osteoblast-like cells).


e5




Table 3. Examples of fish systems to discover molecules affecting skeleton and bone formation Molecules

Fish systems

Skeletal and bone effects

References

2,3,7,8-Tetrachlorodibenzo-p-dioxin (TCDD) 17b-Estradiol 3-Methylcholanthrene

Developing medaka Cultured goldfish scales Developing zebrafish Regenerating zebrafish Cultured gilthead seabream bone cells Cultured goldfish scales Cultured gilthead seabream bone cells Cultured goldfish scales Developing zebrafish

Impaired chondro- and osteogenesis Increased osteoclastic activity Increased rate of skeletal deformities Reduced de novo bone formation Reduced ECM mineralization Suppressed osteoclastic activity Reduced ECM mineralization Suppressed osteoclastic and osteoblastic activites Craniofacial defects

[87] [58] [36]

Developing zebrafish Iron-overload zebrafish Developing zebrafish Developing zebrafish

Increased bone formation Rescued iron-induced osteoporosis Bone loss Reduced bone mineralization

[31] [28] [31] [89]

Calcitonin Vanadium Bisphenol A Ethyl tert butyl ether (ETBE) Tertiary amyl methyl ether (TAME) Vitamin D3 analogs Deferoxamine Parathyroid hormone (PTH) Dorsomorphin

regeneration, that is the complete reformation of missing tissues [33–35]. When caudal fin tissues are amputated, a thick wound epidermis is formed at the amputation plane, then a mass of undifferentiated mesenchymal cells, called blastema, proliferate beneath the wound epidermis and will later differentiate into the various cell types necessary to the faithful restoration of missing fin structures. Full regeneration is usually achieved after 10–15 days depending on the fish species, the level of amputation and culture conditions. Although zebrafish are raised at 28 8C, assays of fin regeneration are typically performed at 33 8C to speed up regeneration process, which is well advanced after only 5 days. Fin regeneration and de novo bone mineralization is determined after alizarin red staining, imaging and morphometric analysis of regenerated areas (Fig. 1d; see brief protocol in [36])

Mineralogenesis in cell and scale cultures In addition and as a complement to in vivo studies, mineralogenic cell lines developed from several fish species [37–39] can be used to study more in depth mechanisms affecting bone and cartilage cell function and metabolism and the molecular pathways involved in various processes such as extracellular matrix mineralization [40–46] or the mineralogenic effect of various molecules, for example retinoic acid [47,48], polyunsaturated fatty acids [49], vanadate [46,50–52] and polycyclic aromatic hydrocarbon [36]. In vitro mineralization is easily detected and quantified through the alizarin red S staining of hydroxyapatite-like crystals deposited within the extracellular matrix of fish bone-derived cell lines upon exposure to a mineralogenic cocktail ([37]; Fig. 1f). While cell cultures have been used to study mechanisms of extracellular matrix mineralization, they have limited value in the study of cell–cell and cell–matrix interactions. Elasmoid scales of teleost fish (Fig. 1e) are dermal bone elements that develop late (30 dpf in zebrafish) and serve as a reservoir of calcium [53,54]. They are small, easily accessible, abundant e6

[58] [52] [57] [88]

(hundreds of highly similar scales per fish), transparent and have the ability to quickly regenerate (4 weeks in zebrafish), when lost or removed [25,55]. As for regenerating fin rays, scales have been recently used in biomedical studies aiming at the better understanding of mechanism of bone regeneration [25,55] but also as an in vivo disease model for osteoporosis studies [56]. The possibility of culturing fish scales in vitro as a bone unit, where osteoclasts and osteoblasts cohabit on both sides of the mineralized matrix and interact in a way resembling in vivo conditions, has allowed their use to assess effects of anti/pro-osteogenic molecules [57,58] and as a valuable ex vivo system to discover new drugs affecting bone formation and/or resorption.

Conclusions Through the availability of mutant and transgenic lines, fish has become an attractive model system to study bone disorders. Furthermore, fish provide unique insights into underlying molecular mechanisms that cannot be gained in traditional mammalian systems due to shortcomings, morphological constrains and technical limitations. Fish are also cost-effective models with the potential to identify, in a shorter time, novel drugs to treat bone disorders as well as screening for osteotoxic molecules. Finding new treatments for human diseases is challenging and future works should aim at developing novel fish systems capable of modeling more bone disorders and at automating screening processes.

Conflict of interest The authors have no conflict of interest to declare.

Acknowledgements This work was supported by the Portuguese Funding Agency for Science and Technology (FCT) through AQUATOX (PTDC/MAR/112992/2009) research project, by the Atlantic Area Transnational Cooperation Programme of the European





Community through MARMED (2011-1/164) project, and by the National Strategic Reference Framework (QREN I&DT) through ZEBRAFEEDS (QREN 23000) project.

[22]

[23]

References [1] Hall BK. Bones and cartilage: developmental and evolutionary skeletal biology. Academic Press; 2005. [2] Javidan Y, Schilling TF. Development of cartilage and bone. Methods Cell Biol 2004;76:415–36. [3] Apschner A, Schulte-Merker S, Witten PE. Not all bones are created equal – using zebrafish and other teleost species in osteogenesis research. Methods Cell Biol 2011;105:239–55. [4] Hall BK, Witten PE. Plasticity of and transitions between skeletal tissues in vertebrate evolution and development. In: Anderson JS, Sues H-DD., editors. Major transitions in vertebrate evolution. Indiana University Press; 2007. p. 13–56. [5] Witten PE, Huysseune A. A comparative view on mechanisms and functions of skeletal remodelling in teleost fish, with special emphasis on osteoclasts and their function. Biol Rev Camb Philos Soc 2009;84: 315–46. [6] Arratia G, Schultze H-P. Reevaluation of the caudal skeleton of certain actinopterygian fishes: III. Salmonidae. Homologization of caudal skeletal structures. J Morphol 1992;249:187–249. [7] Nordvik K, Kryvi H, Totland GK, Grotmol S. The salmon vertebral body develops through mineralization of two preformed tissues that are encompassed by two layers of bone. J Anat 2005;206:103–14. [8] Benjamin M. The cranial cartilages of teleosts and their classification. J Anat 1990;169:153–72. [9] Witten PE, Huysseune A, Hall BK. A practical approach for the identification of the many cartilaginous tissues in teleost fish. J Appl Ichthyol 2010;26:257–62. [10] Lieschke GJ, Currie PD. Animal models of human disease: zebrafish swim into view. Nat Rev Genet 2007;8:353–67. [11] Howe K, Clark MD, Torroja CF, Torrance J, Berthelot C, Muffato M, et al. The zebrafish reference genome sequence and its relationship to the human genome. Nature 2013;496:498–503. [12] Langheinrich U. Zebrafish: a new model on the pharmaceutical catwalk. Bioessays 2003;25:904–12. [13] Spoorendonk KM, Hammond CL, Huitema LFA, Vanoevelen J, SchulteMerker S. Zebrafish as a unique model system in bone research: the power of genetics and in vivo imaging. J Appl Ichthyol 2010;26:219–24. [14] Santoriello C, Zon LI. Hooked! Modeling human disease in zebrafish. J Clin Invest 2012;122:2337–43. [15] Hosen MJ, Vanakker OM, Willaert A, Huysseune A, Coucke P, De Paepe A. Zebrafish models for ectopic mineralization disorders: practical issues from morpholino design to post-injection observations. Front Genet 2013;4:74. [16] Huitema LFA, Apschner A, Logister I, Spoorendonk KM, Bussmann J, Hammond CL, et al. Entpd5 is essential for skeletal mineralization and regulates phosphate homeostasis in zebrafish. Proc Natl Acad Sci U S A 2012;109:21372–77. [17] DeLaurier A, Nakamura Y, Braasch I, Khanna V, Kato H, Wakitani S, et al. Histone deacetylase-4 is required during early cranial neural crest development for generation of the zebrafish palatal skeleton. BMC Dev Biol 2012;12:16. [18] Hammond CL, Moro E. Using transgenic reporters to visualize bone and cartilage signaling during development in vivo. Front Endocrinol (Lausanne) 2012;3:91. [19] Knopf F, Hammond C, Chekuru A, Kurth T, Hans S, Weber CW, et al. Bone regenerates via dedifferentiation of osteoblasts in the zebrafish fin. Dev Cell 2011;20:713–24. [20] Nichols JT, Pan L, Moens CB, Kimmel CB. Barx1 represses joints and promotes cartilage in the craniofacial skeleton. Development 2013;140:2765–75. ¨ ttner A, To TT, Chan SJH, Winkler C. A col10a1:nlGFP [21] Renn J, Bu transgenic line displays putative osteoblast precursors at the

[24]

[25]

[26]

[27]

[28]

[29]

[30] [31]

[32] [33]

[34]

[35] [36] [37]

[38]

[39]

[40]

[41]

[42]

medaka notochordal sheath prior to mineralization. Dev Biol 2013;381:134–43. Brittijn SA, Duivesteijn SJ, Belmamoune M, Bertens LFM, Bitter W, de Bruijn JD, et al. Zebrafish development and regeneration: new tools for biomedical research. Int J Dev Biol 2009;53:835–50. Fleming A. Zebrafish as an alternative model organism for disease modelling and drug discovery: implications for the 3Rs [WWW Document]; 2007, http://www.nc3rs.org.uk/news.asp?id=421. ¨ hr H, Hammerschmidt M. Zebrafish in endocrine systems: recent Lo advances and implications for human disease. Annu Rev Physiol 2011;73:183–211. Metz JR, de Vrieze E, Lock E-J, Schulten IE, Flik G. Elasmoid scales of fishes as model in biomedical bone research. J Appl Ichthyol 2012;28:382–7. Renn J, Winkler C, Schartl M, Fischer R, Goerlich R. Zebrafish and medaka as models for bone research including implications regarding spacerelated issues. Protoplasma 2006;229:209–14. Baraban SC, Dinday MT, Hortopan GA. Drug screening in Scn1a zebrafish mutant identifies clemizole as a potential Dravet syndrome treatment. Nat Commun 2013;4:2410. Chen B, Yan Y-L, Liu C, Bo L, Li G-F, Wang H, et al. Therapeutic effect of deferoxamine on iron overload-induced inhibition of osteogenesis in a zebrafish model. Calcif Tissue Int 2014;94:353–60. Gavaia PJ, Simes DC, Ortiz-Delgado JB, Viegas CSB, Pinto JP, Kelsh RN, et al. Osteocalcin and matrix Gla protein in zebrafish (Danio rerio) and Senegal sole (Solea senegalensis): comparative gene and protein expression during larval development through adulthood. Gene Expr Patterns 2006;6:637–52. Walker MB, Kimmel CB. A two-color acid-free cartilage and bone stain for zebrafish larvae. Biotech Histochem 2007;82:23–8. Fleming A, Sato M, Goldsmith P. High-throughput in vivo screening for bone anabolic compounds with zebrafish. J Biomol Screen 2005;10:823– 31. Mackay EW, Apschner A, Schulte-Merker S. A bone to pick with zebrafish. Bonekey Rep 2013;2:445. Akimenko M-A, Smith A. Paired fin repair and regeneration. In: Hall BK, editor. Fins into limbs: evolution, development, and transformation. The University of Chicago Press; 2007. p. 152–62. Nakatani Y, Kawakami A, Kudo A. Cellular and molecular processes of regeneration, with special emphasis on fish fins. Dev Growth Differ 2007;49:145–54. Yoshinari N, Kawakami A. Mature and juvenile tissue models of regeneration in small fish species. Biol Bull 2011;221:62–78. Laize´ V, Gavaia PJ, Viegas MN, Caria J, Luis N. Osteotoxicity of 3methylcholanthrene in fish. Aquat Procedia 2014 [in press]. Pombinho AR, Laize´ V, Molha DM, Marques SMP, Cancela ML. Development of two bone-derived cell lines from the marine teleost Sparus aurata; evidence for extracellular matrix mineralization and cell-typespecific expression of matrix Gla protein and osteocalcin. Cell Tissue Res 2004;315:393–406. Rafael MS, Marques CL, Parameswaran V, Cancela ML, Laize´ V. Fish bonederived cell lines: an alternative in vitro cell system to study bone biology. J Appl Ichthyol 2010;26:230–4. Vijayakumar P, Laize´ V, Cardeira J, Trindade M, Cancela ML. Development of an in vitro cell system from zebrafish suitable to study bone cell differentiation and extracellular matrix mineralization. Zebrafish 2013;10:500–9. Fonseca VG, Rosa J, Laize´ V, Gavaia PJ, Cancela ML. Identification of a new cartilage-specific S100-like protein up-regulated during endo/ perichondral mineralization in gilthead seabream. Gene Expr Patterns 2011;11:448–55. Fonseca VG, Laize´ V, Valente MS, Cancela ML. Identification of an osteopontin-like protein in fish associated with mineral formation. FEBS J 2007;274:4428–39. Laize´ V, Pombinho AR, Cancela ML. Characterization of Sparus aurata osteonectin cDNA and in silico analysis of protein conserved features: evidence for more than one osteonectin in Salmonidae. Biochimie 2005;87:411–20.


e7



[43] Rafael MS, Laize´ V, Florindo C, Ferraresso S, Bargelloni L, Cancela ML. Overexpression of four and a half LIM domains protein 2 promotes epithelial-mesenchymal transition-like phenotype in fish pre-osteoblasts. Biochimie 2012;94:1128–34. [44] Rafael MS, Laize´ V, Cancela ML. Identification of Sparus aurata bone morphogenetic protein 2: molecular cloning, gene expression and in silico analysis of protein conserved features in vertebrates. Bone 2006;39:1373–81. [45] Tiago DM, Marques CL, Roberto VP, Cancela ML, Laize´ V. Mir-20a regulates in vitro mineralization and BMP signaling pathway by targeting BMP-2 transcript in fish. Arch Biochem Biophys 2014;543:23–30. [46] Tiago DM, Laize´ V, Bargelloni L, Ferraresso S, Romualdi C, Cancela ML. Global analysis of gene expression in mineralizing fish vertebra-derived cell lines: new insights into anti-mineralogenic effect of vanadate. BMC Genomics 2011;12:310. ˜ es B, Pombinho AR, Cancela ML. Retinoic acid [47] Conceiçaõ N, Laize´ V, Simo is a negative regulator of matrix Gla protein gene expression in teleost fish Sparus aurata. Biochim Biophys Acta 2008;1779:28–39. [48] Fernańdez I, Tiago DM, Laize´ V, Cancela ML, Gisbert E. Retinoic acid differentially affects in vitro proliferation, differentiation and mineralization of two fish bone-derived cell lines: different gene expression of nuclear receptors and ECM proteins. J Steroid Biochem Mol Biol 2014;140:34–43. [49] Viegas MN, Dias J, Cancela ML, Laize´ V. Polyunsaturated fatty acids regulate cell proliferation, extracellular matrix mineralization and gene expression in a gilthead seabream skeletal cell line. J Appl Ichthyol 2012;28:427–32. [50] Tiago DM, Cancela ML, Laize´ V. Proliferative and mineralogenic effects of insulin, IGF-1, and vanadate in fish osteoblast-like cells. J Bone Miner Metab 2011;29:377–82. [51] Tiago DM, Cancela ML, Aureliano M, Laize´ V. Vanadate proliferative and anti-mineralogenic effects are mediated by MAPK and PI-3K/Ras/Erk pathways in a fish chondrocyte cell line. FEBS Lett 2008;582: 1381–5. [52] Tiago DM, Laize´ V, Cancela ML, Aureliano M. Impairment of mineralization by metavanadate and decavanadate solutions in a fish bone-derived cell line. Cell Biol Toxicol 2008;24:253–63. [53] Carragher JF, Sumpter JP. The mobilization of calcium from calcified tissues of rainbow trout (Oncorhynchus mykiss) induced to synthesize vitellogenin. Comp Biochem Physiol Part A Physiol 1991;99: 169–72. [54] Mugiya Y, Watabe N. Studies on fish scale formation and resorption II: effect of estradiol on calcium homeostasis and skeletal tissue resorption in the goldfish, Carassius auratus, and the killifish, Fundulus heteroclitus. Comp Biochem Physiol Part A Physiol 1977;57:197–202. [55] De Vrieze E, Sharif F, Metz JR, Flik G, Richardson MK. Matrix metalloproteinases in osteoclasts of ontogenetic and regenerating zebrafish scales. Bone 2011;48:704–12. [56] De Vrieze E, van Kessel MAHJ, Peters HM, Spanings FAT, Flik G, Metz JR. Prednisolone induces osteoporosis-like phenotype in regenerating zebrafish scales. Osteoporos Int 2014;25:567–78. [57] Suzuki N, Hattori A. Bisphenol A suppresses osteoclastic and osteoblastic activities in the cultured scales of goldfish. Life Sci 2003;73:2237–47. [58] Suzuki N, Suzuki T, Kurokawa T. Suppression of osteoclastic activities by calcitonin in the scales of goldfish (freshwater teleost) and nibbler fish (seawater teleost). Peptides 2000;21:115–24. [59] Shahar R, Dean MN. The enigmas of bone without osteocytes. Bonekey Rep 2013;2:343. [60] Cao L, Moriishi T, Miyazaki T, Iimura T, Hamagaki M, Nakane A, et al. Comparative morphology of the osteocyte lacunocanalicular system in various vertebrates. J Bone Miner Metab 2011;29:662–70. [61] Totland GK, Fjelldal PG, Kryvi H, Løkka G, Wargelius A, Sagstad A, et al. Sustained swimming increases the mineral content and osteocyte density of salmon vertebral bone. J Anat 2011;219:490–501. [62] Turner CH, Warden SJ, Bellido T, Plotkin LI, Kumar N, Jasiuk I, et al. Mechanobiology of the skeleton. Sci Signal 2009;2:pt3.

e8


[63] Witten PE, Hansen A, Hall BK. Features of mono- and multinucleated bone resorbing cells of the zebrafish Danio rerio and their contribution to skeletal development, remodeling, and growth. J Morphol 2001;250:197– 207. [64] You L, Temiyasathit S, Lee P, Kim CH, Tummala P, Yao W, et al. Osteocytes as mechanosensors in the inhibition of bone resorption due to mechanical loading. Bone 2008;42:172–9. [65] Fisher S, Jagadeeswaran P, Halpern ME. Radiographic analysis of zebrafish skeletal defects. Dev Biol 2003;264:64–76. [66] Asharani PV, Keupp K, Semler O, Wang W, Li Y, Thiele H, et al. Attenuated BMP1 function compromises osteogenesis, leading to bone fragility in humans and zebrafish. Am J Hum Genet 2012;90:661–74. [67] To TT, Witten PE, Renn J, Bhattacharya D, Huysseune A, Winkler C. Ranklinduced osteoclastogenesis leads to loss of mineralization in a medaka osteoporosis model. Development 2012;139:141–50. [68] Barrett R, Chappell C, Quick M, Fleming A. A rapid, high content, in vivo model of glucocorticoid-induced osteoporosis. Biotechnol J 2006;1: 651–5. [69] Eames BF, Yan Y-L, Swartz ME, Levic DS, Knapik EW, Postlethwait JH, et al. Mutations in fam20b and xylt1 reveal that cartilage matrix controls timing of endochondral ossification by inhibiting chondrocyte maturation. PLoS Genet 2011;7:e1002246. [70] Cle´ment A, Wiweger M, von der Hardt S, Rusch MA, Selleck SB, Chien C-B, et al. Regulation of zebrafish skeletogenesis by ext2/dackel and papst1/ pinscher. PLoS Genet 2008;4:e1000136. [71] Wiweger MI, Zhao Z, van Merkesteyn RJP, Roehl HH, Hogendoorn PCW. HSPG-deficient zebrafish uncovers dental aspect of multiple osteochondromas. PLoS One 2012;7:e29734. [72] Flanagan-Steet H, Sias C, Steet R. Altered chondrocyte differentiation and extracellular matrix homeostasis in a zebrafish model for mucolipidosis II. Am J Pathol 2009;175:2063–75. [73] Laue K, Pogoda H-M, Daniel PB, van Haeringen A, Alanay Y, von Ameln S, et al. Craniosynostosis and multiple skeletal anomalies in humans and zebrafish result from a defect in the localized degradation of retinoic acid. Am J Hum Genet 2011;89:595–606. [74] Spoorendonk KM, Peterson-Maduro J, Renn J, Trowe T, Kranenbarg S, Winkler C, et al. Retinoic acid and Cyp26b1 are critical regulators of osteogenesis in the axial skeleton. Development 2008;135: 3765–74. [75] Bauer H, Lele Z, Rauch GJ, Geisler R, Hammerschmidt M. The type I serine/ threonine kinase receptor Alk8/Lost-a-fin is required for Bmp2b/7 signal transduction during dorsoventral patterning of the zebrafish embryo. Development 2001;128:849–58. [76] Gorman KF, Tredwell SJ, Breden F. The mutant guppy syndrome curveback as a model for human heritable spinal curvature. Spine (Phila Pa 1976) 2007;32:735–41. [77] Schauerte HE, van Eeden FJM, Fricke C, Odenthal J, Stra¨hle U, Haffter P. Sonic hedgehog is not required for the induction of medial floor plate cells in the zebrafish. Development 1998;125:2983–93. [78] Yan Y, Miller CT, Nissen RM, Singer A, Liu D, Kirn A, et al. A zebrafish sox9 gene required for cartilage morphogenesis. Development 2002;129:5065–79. [79] Nissen RM, Amsterdam A, Hopkins N. A zebrafish screen for craniofacial mutants identifies wdr68 as a highly conserved gene required for endothelin-1 expression. BMC Dev Biol 2006;6:28. [80] Piotrowski T, Ahn D, Schilling TF, Nair S, Ruvinsky I, Geisler R, et al. The zebrafish van Gogh mutation disrupts tbx1, which is involved in the DiGeorge deletion syndrome in humans. Development 2003;130:5043–52. [81] Lang MR, Lapierre LA, Frotscher M, Goldenring JR, Knapik EW. Secretory COPII coat component Sec23a is essential for craniofacial chondrocyte maturation. Nat Genet 2006;38:1198–203. [82] Melville DB, Montero-Balaguer M, Levic DS, Bradley K, Smith JR, Hatzopoulos AK, et al. The feelgood mutation in zebrafish dysregulates COPII-dependent secretion of select extracellular matrix proteins in skeletal morphogenesis. Dis Model Mech 2011;4:763–76.





[83] Eames BF, Singer A, Smith GA, Wood ZA, Yan Y-L, He X, et al. UDP xylose synthase 1 is required for morphogenesis and histogenesis of the craniofacial skeleton. Dev Biol 2010;341:400–15. [84] Sarmah S, Barrallo-Gimeno A, Melville DB, Topczewski J, Solnica-Krezel L, Knapik EW. Sec24D-dependent transport of extracellular matrix proteins is required for zebrafish skeletal morphogenesis. PLoS ONE 2010;5:e10367. [85] Xi Y, Chen D, Sun L, Li Y, Li L. Characterization of zebrafish mutants with defects in bone calcification during development. Biochem Biophys Res Commun 2013;440:132–6. [86] Green J, Taylor JJ, Hindes A, Johnson SL, Goldsmith MI. A gain of function mutation causing skeletal overgrowth in the rapunzel mutant. Dev Biol 2009;334:224–34.

[87] Dong W, Hinton DE, Kullman SW. TCDD disrupts hypural skeletogenesis during medaka embryonic development. Toxicol Sci 2012;125:91–104. [88] Bonventre JA, White LA, Cooper KR. Craniofacial abnormalities and altered wnt and mmp mRNA expression in zebrafish embryos exposed to gasoline oxygenates ETBE and TAME. Aquat Toxicol 2012;120– 121:45–53. [89] Yu PB, Hong CC, Sachidanandan C, Babitt JL, Deng DY, Hoyng SA, et al. Dorsomorphin inhibits BMP signals required for embryogenesis and iron metabolism. Nat Chem Biol 2008;4:33–41. [90] Neuhauss SCF, Solnica-Krezel L, Schier AF, Zwartkruis F, Stemple DL, Malicki J, et al. Mutations affecting craniofacial development in zebrafish. Development 1996;123:357–67.


e9
