PRIMER
Model systems for regeneration: zebrafish
Ines J. Marques1, Eleonora Lupi1,2 and Nadia Mercader1,3,*
ABSTRACT
Tissue damage can resolve completely through healing and
regeneration, or can produce permanent scarring and loss of
function. The response to tissue damage varies across tissues and
between species. Determining the natural mechanisms behind
regeneration in model organisms that regenerate well can help us
develop strategies for tissue recovery in species with poor regenerative
capacity (such as humans). The zebrafish (Danio rerio) is one of the
most accessible vertebratemodels to study regeneration. In this Primer,
we highlight the tools available to study regeneration in the zebrafish,
provide an overview of the mechanisms underlying regeneration in this
system and discuss future perspectives for the field.
KEY WORDS: Regeneration, Zebrafish, Cell progenitors, Injury
methods, Genetic tools
Introduction
Most vertebrate tissues undergo continuous cell turnover to
maintain tissue homeostasis, and cells that no longer exert their
function correctly are eliminated and replaced by new ones. This
continuous cellular turnover is not homogenous in all tissues:
whereas gut enterocytes and skin keratinocytes show a high turnover
(Young and Heath, 2000), cardiomyocytes (Bergmann et al., 2009;
Senyo et al., 2013) and neurons are rarely replaced throughout adult
life. In the case of neurons, proliferation is limited to specific
regions in the brain (Ernst et al., 2014; Lim and Alvarez-Buylla,
2016). Beyond homeostatic tissue maintenance, some tissues and
organs are capable of reacting to an injury by replacing the lesioned
region. In this scenario, responses can either involve tissue repair,
where a fibrotic scar replaces the injury, or regeneration, where new
cells proliferate and/or differentiate into the same cell type as the one
that was damaged, thus reconstituting the tissue so that it can recover
its functional activity. As a general outline, during injury response
the damaged area (Fig. 1A,B) is initially infiltrated by immune cells
(Fig. 1C), which clean the dead tissue and prevent infections. This is
followed by the deposition of extracellular matrix (ECM) (Fig. 1D),
which provides support and structure to the injured area. ECM
deposition results in the development of a, sometimes permanent,
fibrotic scar. Alternatively, the lesioned region can be replaced
by new tissue (Fig. 1E) in a multi-step regenerative process that
can involve dedifferentiation, proliferation and re-differentiation
of existing cells (as discussed further below). In some cases, the
regenerative phase occurs concomitant with the regression of the
transient fibrotic tissue. Overall, the resulting regenerated tissue
has the same morphological and physiological properties as the
lost tissue, and is hence fully functional (Fig. 1F) (Hardy, 1989).
Vertebrates vary widely in their regenerative capacity. Whereas
mammals usually have a more limited regenerative capacity –
restricted to few tissues or organs (e.g. skin and liver) or a short time
window after birth [e.g. the heart (Porrello et al., 2011)] – other
vertebrates possess a higher degree of regenerative capacity: e.g.
axolotls are capable of regenerating several organs, including the
heart (Cano-Martinez et al., 2010) and the limbs (Simon and Tanaka,
2013). Among the vertebrate animal models available for
regeneration studies, the zebrafish (Danio rerio) – a 2-5 cm long
freshwater teleost – is one of the most widely used. Following their
establishment as a genetically and experimentally accessible model
system for studying development (Haffter et al., 1996; Streisinger
et al., 1981), early regeneration studies showed that zebrafish, like
many other fish, are able to fully regenerate many tissues and organs
(Bernhardt et al., 1996; Johnson andWeston, 1995; Poss et al., 2002;
Raya et al., 2003; Rowlerson et al., 1997; White et al., 1994). This
pioneering research convinced the scientific community to consider
this model organism for studying organ regeneration as well as
development. Some of the tissues and organs most typically used in
regeneration studies include the spinal cord (Becker et al., 1998),
brain and cerebellum (Kroehne et al., 2011; Liu et al., 2004), retina
(Vihtelic and Hyde, 2000), hair cells (Harris et al., 2003), heart (Poss
et al., 2002; Raya et al., 2003), caudal fin (Nechiporuk and Keating,
2002; White et al., 1994), kidney (Reimschuessel, 2001), and liver
(Burkhardt-Holm et al., 1999).
Regeneration studies have been greatly facilitated by the ever-
increasing power of genetic tools, including transgenesis and
genome editing to generate loss-of-function mutations. (Hwang
et al., 2013; Kawakami, 2007; Moore et al., 2012; Stuart et al.,
1988). In comparison with other highly regenerative animal models
such as the axolotl, these were developed much earlier in zebrafish,
Model systems for regeneration
This article is part of a series entitled ‘Model systems for regeneration’.
This series of articles aims to highlight key model systems and species
that are currently being used to study tissue and organ regeneration.
Each article provides background information about the phylogenetic
position of the species, its life-cycle and habitat, the different organs and
tissues that regenerate, and the experimental tools and techniques that
are available for studying these organisms in a regenerative context.
Importantly, these articles also give examples of how the study of these
models has increased our understanding of regenerative mechanisms
more broadly, and how some of the open questions in the field of
regeneration may be answered using these organisms. To see the full
collection as it grows, please visit: https://dev.biologists.org/collection/
regeneration_models
Received 13 February 2019; Accepted 19 August 2019
1Institute of Anatomy, University of Bern, Bern 3012, Switzerland. 2Acquifer, Ditabis,
Digital Biomedical Imaging Systems, Pforzheim, Germany. 3Centro Nacional de
Investigaciones Cardiovasculares CNIC, Madrid 2029, Spain.
*Author for correspondence (nadia.mercader@ana.unibe.ch)
I.J.M., 0000-0003-1254-9394; E.L., 0000-0003-0509-3616; N.M., 0000-0002-
0905-6399
This is an Open Access article distributed under the terms of the Creative Commons Attribution
License (https://creativecommons.org/licenses/by/4.0), which permits unrestricted use,
distribution and reproduction in any medium provided that the original work is properly attributed.
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© 2019. Published by The Company of Biologists Ltd | Development (2019) 146, dev167692. doi:10.1242/dev.167692
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further stimulating its use for regeneration studies. Moreover,
excellent platforms for sharing resources and knowledge
(summarized in Table 1) are contributing to a steady growth of
the research community working with this model organism, also in
the context of regeneration. In this Primer, we provide a broad
overview of the available tools for regeneration research in the
zebrafish, and focus on the challenges and opportunities presented
by this model organism.
Tools for studying regeneration in zebrafish
Many of the attributes that make zebrafish an appealing model for
developmental biologists also give it significant advantages for
regeneration researchers – with the genetic and genomic tools, and
accessibility for live imaging and drug screening being particularly
important. We discuss these topics further below, but focus first on
the range of injury models available to researchers.
Tissue injury models
Several injury models have been established to study regeneration in
zebrafish, including physical injury models such as resection,
stabbing or cryoinjury, or, alternatively, genetic procedures to ablate
a specific cell type (Table 2 and Fig. 2). Although most studies are
performed in adult zebrafish, in some instances larvae are also used
to study organ regeneration. Choosing the appropriate injury model,
and developmental stage, is an important decision. First, one should
take into account the type of injury intended to be mimicked –
particularly if the primary aim is to provide a model for a human
condition. However, it is not always possible to mimic a mammalian
injury process. For example, myocardial infarction is caused by an
ischemic event, often due to the occlusion of coronary vessels by
an atherosclerotic plaque. This type of injury can be mimicked in
mice, but is very challenging to recapitulate in the zebrafish, owing
to its small size. Second, the choice of the injury method also
depends on the biological question to be answered. To study global
processes of regeneration, one probably needs to use a method that
lesions the whole tissue, such as surgically induced injuries. If the
aim is to understand the regenerative capacity of a specific cell type,
then genetic ablation might be a better choice. As a further
consideration, different types of injury can lead to changes in the
efficiency of cell replacement or speed of regeneration, as
Table 1. Online resources for the zebrafish community
Resource Description URL and references
Zebrafish Information Network
(ZFIN)
Database of multiple resources available for zebrafish studies,
including protocols and available lines
http://www.zfin.org (Howe et al., 2013)
International Zebrafish Society
(IZFS)
Promotion of zebrafish research, organization of conferences,
travel fellowships and awards
https://www.izfs.org/
EUFishBiomed Promotes exchange of information within the fish community,
support of conferences and travel fellowships
https://www.eufishbiomed.kit.edu/ (Strahle et al., 2012)
Zebrafish International
Resource Center (ZIRC)
Zebrafish stock center (USA) collecting existing zebrafish
resources
https://zebrafish.org/
Karlsruhe Institute of
Technology (KIT)
Zebrafish stock center (Europe) collecting existing zebrafish
resources and providing training
https://www.ezrc.kit.edu/ (Geisler et al., 2016)
China Zebrafish Resource
Center (CZRC)
China Zebrafish Resource Center collecting existing zebrafish
resources, developing new lines and technology
http://en.zfish.cn/
Tissue cell FibroblastImmune cell
Platelet ECM Blood vessel
A B C
D E F
Injured cells
Proliferating cell
Key
Fig. 1. Phases of tissue regeneration. (A) Uninjured tissue. (B) Injury: injured area is shaded in gray, representing damaged and dead cells. The blood
vessel in direct contact with the lesion is also damaged and is marked in a lighter red color than the healthy vessels. (C) Wound closure and immune response:
the injured area is infiltrated by platelets to stop the bleeding resulting from injury to the blood vessels in the damaged area. At the same time, the lesion
is also infiltrated by immune cells. (D) Clearing of debris and deposition of extracellular matrix (ECM): the immune cells start to clean the dead tissue and
other debris resulting from the injury. Fibroblasts proliferate and produce ECM fibers that fill the cleared injury area to prevent the injury from collapsing.
The formation of new blood vessels takes place (thinner red lines). (E) Resorption of ECM and cell proliferation: the ECM starts to be eliminated as the injury is
repopulated by new cells through proliferation. (F) Regeneration: the injured area has been completely repopulated and the tissue reconstituted to its
homeostatic condition. Cells undergo cell cycle arrest and very few proliferating cells can be seen.
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exemplified for the spinal cord (Ohnmacht et al., 2016) and the heart
(Gonzalez-Rosa et al., 2011).
Surgical injuries
One of themost common surgicalmethods is resection (amputation) of
the tissue. This technique is commonly used for fin and heart.
Following fin amputation, the first step towards regeneration involves
the development of an epidermal cap that closes the injury.Underneath,
a blastema comprising undifferentiated cells is formed (discussed
further below). After initial proliferation, these cells redifferentiate,
culminating in the formation of new fin tissue (reviewed by Pfefferli
and Jazẃinśka, 2015). In the heart, the first step towards regeneration
after ventricular resection is the formation of a blood clot, later replaced
by a fibrin clot, followed by de novo growth to replace the lost tissue,
until the damagedmyocardiumbecomes almost indistinguishable from
the surrounding tissue (reviewed by Gonzalez-Rosa et al., 2017).
Another popular surgically induced lesion method is cryoinjury,
which is frequently used for the heart (Chablais et al., 2011;
Gonzalez-Rosa et al., 2011; Schnabel et al., 2011) and has also been
described in fin (Chassot et al., 2016). In this method, the tissue is
exposed to a super-cooled metal, which effectively kills the cells it
touches. In the fin, after the dead tissue falls off, regeneration
proceeds as in the amputation model (Chassot et al., 2016). By
contrast, the regenerative process in the heart is a little different:
it begins with the replacement of the necrotic tissue with
ECM-depositing fibroblasts. Subsequently, the transient scar
tissue regresses and is replaced by new cardiac tissue (Gonzalez-
Rosa et al., 2012). As well as low temperatures, cell death can be
induced by exposure to high temperatures, either using a heated
metal rod (Raymond et al., 2006) or by applying a light source in a
determined region (Conedera et al., 2017; Vihtelic and Hyde, 2000).
Finally, another method is stabbing, commonly used to damage
the central nervous system, including the spinal cord (Becker et al.,
1998). In this case, accumulation and proliferation of glial cells in
the damaged area is followed by the proliferation of progenitors that
ultimately differentiate into new nervous tissue (März et al., 2011).
Chemically induced cell damage
Besides physical stimuli, organ injuries can be induced by chemical
compounds, which can either be directly administered to each
individual fish by intraperitoneal injection, perorally or by dissolving
in the fish water followed by ingestion. The toxicity of gentamicin is
used to produce kidney or lateral line injuries (Kamei et al., 2015) and
acetaminophen is used to induce liver injuries (Cox et al., 2014). After
an initial phase of cell death, proliferation and cell migration lead to the
repopulation of the damaged area, as has been demonstrated in studies
on zebrafish kidney (Reimschuessel and Williams, 1995), lateral line
hair cell (Harris et al., 2003) and liver (Burkhardt-Holm et al., 1999)
regeneration. This method, however, presents some disadvantages,
such as off-target effects, as well as a less uniform injuries caused by
unequal drug distribution (Strahle and Grabher, 2010).
Genetic ablation
This approach allows tissue- or cell type-specific death and can also
provide temporal control of cell ablation. In several transgenic zebrafish
lines, cell death can be specifically induced in targeted cells. Methods
include the regulated expression of diphtheria toxin A chain (Wang
et al., 2011) and overexpression of a cytotoxic fluorescent protein,
KillerRed – which has been used to ablate specific cell types through
photoxicity upon intense light illumination (Del Bene et al., 2010).
Perhaps the most widely used system is based on the ability of the
Table 2. Injury models for zebrafish organ regeneration
Organ Injury method References
Fin Resection Geraudie et al. (1993); Pfefferli and Jaźwińska (2015)
Cryoinjury Chassot et al. (2016)
Heart Resection Poss et al. (2002)
Cryoinjury Chablais et al. (2011); Gonzalez-Rosa et al. (2011); Schnabel et al. (2011)
Genetic ablation Wang et al. (2011)
Central nervous system (brain, spinal cord and eye) Nerve crushing Becker and Becker (2008); Bernhardt et al. (1996)
Stabbing März et al. (2011)
Transection Becker and Becker (2008); Becker et al. (1998)
Visible light Vihtelic and Hyde (2000)
Heat Raymond et al. (2006)
Chemical Fimbel et al. (2007)
Pancreas Genetic ablation Pisharath et al. (2007)
Liver Resection Sadler et al. (2007)
Chemical Cox et al. (2014)
Kidney Chemical Reimschuessel (2001)
Sensory hair cells Chemical Harris et al. (2003)
Skin Laser Richardson et al. (2013)
Heart
Liver Pancreas
Retina
Fin
Brain
Spinal cord
Skin
Kidney
Light/heat
Resection
Cryoinjury
Genetic ablation
Resection
CryoinjuryLight/heatChemicalResection
Chemical
Genetic ablation
Chemical
Stabbing/
transection
Stabbing/transection
Fig. 2. Organ regeneration in the zebrafish. Summary of some of the
organs and tissues used for regeneration studies in zebrafish. Preferred
injury models are annotated for each organ.
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bacterial enzyme nitroreductase to convert the prodrug metronidazol
into a cytotoxin, leading to the death of nitroreductase-expressing cells.
This technique was first described in studies on pancreatic β-cell
regeneration in zebrafish (Curado et al., 2007; Pisharath et al., 2007).
Recently, a more potent prodrug, nifurpirinol, has been identified as a
more efficient and specific compound (Bergemann et al., 2018). In all
cases, genetically targeted cells undergo apoptosis. However, in
contrast to the cryoinjury model, ECM deposition is limited (Wang
et al., 2011). Following cell death, cells from the surrounding healthy
tissue begin proliferating to replace the lost tissue.
Genetic tools
As in other fields of zebrafish research, the use of genetic tools is
essential to understand the role played by different genes during
regeneration. In Table 3 we summarize most of the genetic
modification approaches discussed in the ensuing section and
include the main advantages and disadvantages of each of them.
Transgenesis
There is a wide range of tools available for the generation of
transgenic lines. The most commonly used include the use of Tol2
transposons (Kawakami et al., 2000) and I-SceI meganuclease
(Thermes et al., 2002), which facilitate the integration of foreign
DNA into the fish genome. Alternatively, BAC recombineering
(Bussmann and Schulte-Merker, 2011; Suster et al., 2011) allows
the insertion of larger fragments of DNA that often better
recapitulate endogenous gene expression patterns. Of note,
insertion into the genome is non-directed in these approaches.
The lack of a ROSA26-like landing site has been a caveat; the use of
Phi31 Integrase transgenesis (Mosimann et al., 2013) has been
reported as a potential way to address this, and the problem will
hopefully be solved using CRISPR/Cas9 approaches. Transgenic
fluorescent reporter lines used to tag/track a particular cell type or
report a gene expression pattern have proven to be essential for
investigating regeneration.
Inducible gene expression and genetic fate mapping
Different techniques have been established that allow the use of
genetic tools in a spatially and/or temporally controlled manner
during regeneration.
Heat shock
Inducible gene expression can be achieved by using a heat-shock
driven system (Ádám et al., 2000). The heat-shock promoter hsp70
is used to drive expression of the gene of interest. By maintaining
fish at 35-39°C instead of 28°C, gene expression can be activated.
Gal4/UAS
One of the most widely used systems is the Gal4/UAS
transactivation system, which was adapted from Drosophila to the
Table 3. Genetic tools for zebrafish regeneration studies
Method Application Pros Cons
Fluorescent reporter
lines
Cell/tissue imaging Allow evaluation of gene expression pattern by
promoter/enhancer-driven fluorescent
reporter expression
Detection limited to promoter activity, not suited for
fate mapping
Regulatory region might not fully recapitulate
endogenous gene expression pattern
Fluorescent protein expression might be more stable
than the endogenous gene
Photoconvertible
proteins (e.g. KAEDE)
Lineage tracing Activated by light
Produces a faster response than chemically
induced systems
Difficult to apply in non-superficial adult tissues
Transient labelling
Leakiness
CreERT2/loxP Lineage tracing
Gene overexpression
Temporal and spatial conditional
gene expression
Often used for lineage tracing studies
Cannot be turned off
Requires chemical treatment
Can be leaky (Cre activity even without tamoxifen
induction)
Dre/rox Lineage tracing
Gene overexpression
Conditional gene expression
If combined with CreERT2/loxP, it allows
binary recombination studies
Dre can be linked to estrogen or progesterone
receptor to make it inducible (as with Cre)
Cannot be turned off
Not well established in the zebrafish
Fewer lines available than for Cre/Lox
TetOn/TetOff Gene overexpression Specific gene visualization and/or
overexpression in a spatio-temporally
controlled manner
System can be turned on and off as needed
Few available lines
Doxycyclin toxicity not well assessed
Heat-shock promoter Gene overexpression Temperature-dependent inducible expression
Does not require chemical treatment
Gene expression levels are temperature dependent:
precise temperature settings are critical for the
method to work
Promotor not equally efficient in all cell types
Gal4/UAS Gene overexpression
Cell/tissue imaging
Spatially controlled gene expression Transcriptional silencing when Gal4 is expressed
from very active promotors: mostly limits its use to
developmental stages
TALEN Genome editing and
generation of
mutant lines
Highly specific and efficient
More versatile for sequence recognition
than CRISPR
Time consuming
Has not been applied to knock-in approaches
CRISPR/Cas9 Genome editing and
generation of
mutant lines
sgRNA easier to produce than TALEN
Targeting of various genomic regions with
one injection
Off-target effect possible
Use for site-directed insertions still not fully
established
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zebrafish. The system requires the combination of two separate
transgenic lines – one expressing the Gal4 protein in a specific cell
type and the other containing a construct in which a reporter (e.g.
fluorescent protein) or a gene of interest is placed downstream of the
UAS sequence, to which the Gal4 protein binds. When these lines
are crossed, offspring are generated in which the UAS-driven gene
is expressed in those cells expressing Gal4 (reviewed in greater
detail by Halpern et al., 2008). The Gal4 system can also be used in
an inducible manner, by fusing Gal4 to an ERT2 domain, so that
Gal4 is expressed upon the administration of tamoxifen. This
approach has been useful to study hair cell regeneration (Pinto-
Teixeira et al., 2015). However, the Gal4/UAS system in zebrafish is
prone to transcriptional silencing and accumulation of methylation
(Akitake et al., 2011) over time, thus precluding its use in most cases
to study regeneration in adults.
Cre/Lox
A further genetic tool for spatially controlled gene expression is the
Cre/Lox recombination system (Felker and Mosimann, 2016; Hans
et al., 2009; Langenau et al., 2005). It can also be used for lineage
tracing, to assess the origin and fate of cell types during organ
regeneration (Kikuchi et al., 2010). A number of systems allow
temporal control – most commonly, through fusion of Cre to ERT2
(Feil et al., 1997) or by placing Cre under the control of the hsp70
promoter for heat-shock control (Hesselson et al., 2009). This
system has been used for temporal gene expression control during
the analysis of adult heart regeneration (González-Rosa et al., 2018).
It is also possible to use Cre-based approaches to achieve both
spatial and temporal control (Kirchgeorg et al., 2018).
TetOn/TetOff
This system allows temporally defined tissue-specific gene
expression (Wehner et al., 2015). Its main advantage is that gene
expression can be turned on and off by administration or removal of
doxycycline – i.e. highly controlled transient expression is possible.
It has been used to study Wnt signaling during fin and spinal cord
regeneration (Wehner et al., 2014, 2017).
Photoconvertible fluorescent proteins
Finally, photoconvertible Kaede lines have proven useful when the
aim is to trace a specific cell or cluster of cells during regeneration.
This fluorescent protein undergoes conversion, from green to red
light emission, when irradiated under a specific wave-length (Ando
et al., 2002). This method has been used for in vivo tracing studies of
cardiomyocytes during regeneration (Itou et al., 2012). Other
photoconvertible proteins are also available.
Gene knock-down/knockout tools
Formany decades, the analysis of gene function in the zebrafish relied
on non-directed mutagenesis approaches followed by mapping of the
mutation. This was followed by the use of transgenic gain-of-function
studies or the use of dominant-negative forms or morpholino-based
knockdown approaches. Morpholinos are traditionally injected at the
1-cell stage and so cannot easily be used to study regeneration.
However, vivo-Morpholinos have been developed (by GeneTools)
that can be injected into the blood stream or electroporated into a
specific tissue and pass through cell membranes. However, there are
only few reports on their use and even fewer in the context of
regeneration (Bednarek et al., 2015; Hyde et al., 2012; Kizil and
Brand, 2011; Thummel et al., 2006). A better approach to understand
the effect of the downregulation of a specific gene relies on the use of
precise targeted gene editing tools such as TALENs and CRISPR/
Cas9. A better approach to understanding the effect of the
downregulation of a specific gene relies on the use of precise
targeted gene editing tools such as TALENs and CRISPR/Cas9.
These have proven useful for generating null mutants in zebrafish
(Hwang et al., 2013; Jao et al., 2013) and for assessing gene function
during regeneration (Dupret et al., 2017; Dong et al., 2019; Nauroy
et al., 2019; Unal Eroglu et al., 2018). The generation of conditional
knockouts has also recently been achieved by the directed insertion of
loxP sites using CRISPR/Cas9 (Burg et al., 2018). This new approach
will be extremely valuable to address gene function during
regeneration. Several publications compare the use of all these
genetic engineering methods and can provide researchers with the
necessary support to choose the best method for their studies
(Chatterjee and Lufkin, 2011; Rafferty and Quinn, 2018). Although
not yet well established, the possibility of using modified Cas9
versions for transcriptional modulation or induction of epigenetic
modifications is also being explored in the zebrafish (e.g. Liu et al.,
2019; Long et al., 2015).
Importantly, several screening studies, such as those from the
Kawakami (Kawakami et al., 2010) and Brand (Jungke et al., 2013)
labs, have led to the generation of large databases with numerous
fish lines expressing Gal4 or Cre under specific promoters, which
are available to the whole community. These approaches have been
recently reviewed (Albadri et al., 2017; Carney and Mosimann,
2018). Similarly, large open source repositories for zebrafish
mutants are available (Table 1).
Omics approaches
The different ‘omics tools’ that have become available in recent years
are an important asset for helping researchers gain a deeper knowledge
into the mechanisms regulating regeneration. Regarding genomic
tools, ChIPseq (Kang et al., 2016; Xiao et al., 2016), Tomo-Seq (Wu
et al., 2016), single cell RNAseq (Honkoop et al., 2018 preprint) and
histone profiling (Goldman et al., 2017) have been successfully applied
to study regeneration in the zebrafish. RNAseq data have provided
information, for example, on genes involved in axonal regrowth
following spinal cord injury (Mokalled et al., 2016). Complementing
these approaches, proteomics (Saxena et al., 2012) and metabolomics
(Rabinowitz et al., 2017) are now being used in the context of zebrafish
organ regeneration. The possibility of sequencing-based fate mapping
will significantly contribute to the study of organ regeneration
(Spanjaard et al., 2018). Among the current limitations are the
amount of tissue that can be selected for these approaches, annotations
of the zebrafish genome and bioinformatics analysis platforms mainly
oriented to mammalian models.
Live imaging
One of the main advantages of the zebrafish is the possibility of
imaging morphogenetic processes. During embryogenesis, the use
of fluorescent reporter lines facilitates imaging of tissues and organs
at the cellular and subcellular level. Microscopic techniques such as
light sheet microscopy allow for prolonged imaging with minimal
interference of physiological processes that occur during either
regeneration or development (Ding et al., 2018; Power and Huisken,
2017). The zebrafish larva is also sufficiently transparent that live
imaging can be used to follow regenerative processes in organs and
tissues in up to 5-day-old larvae or beyond, depending on the organ.
The preferred injury model during these stages is genetic ablation,
being particularly well suited given the small size of the animals at
this age. Automated systems for larval imaging are being developed
that facilitate the acquisition and analysis of data, and allow high-
throughput screening assays (Early et al., 2018; Pandey et al., 2019).
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In the case of adult zebrafish, in vivo imaging of regenerating
internal organs can prove more difficult. Nevertheless, short- and
long-term imaging of regeneration of external structures have been
achieved using fluorescent light microscopy, as described for caudal
fin neoangiogenesis (Xu et al., 2015), regeneration of the skin
epidermis (Chen et al., 2016) or scale regeneration (Cox et al.,
2018). Although more challenging, even internal structures can
be imaged and followed through time, as has been shown for
adult neural stem cells in homeostasis and during regeneration
(Barbosa et al., 2015; Dray et al., 2015). New fish transgenic
lines have been developed in the hope of improving in vivo
imaging of regenerative processes in the adult. Luciferase-based
transgenic lines have proven useful for regeneration studies,
particularly when looking at internal regenerating structures, as
luciferase bioluminescence can be detected even in deep tissues
(Chen et al., 2013).
Ultrasound or magnetic resonance imaging have also been used
for studying functional recovery after lesion-induced damage
(Goessling et al., 2007; González-Rosa et al., 2014; Hein et al.,
2015; Koth et al., 2017).
Small molecules and drug screening
Zebrafish are also particularly appropriate for small-molecule and
drug-screening studies (Mathew et al., 2007; Matsuda et al., 2018).
In this regard, approaches using genetic ablation injuries in larvae
are most valuable, as a large number of animals can be screened
simultaneously. By using injured zebrafish larvae, multiple
compounds can be tested at the same time and used to identify
those with a pro-regenerative effect and those that would work to
impair regeneration. Screens have identified small molecules with
the potential to induce β-cell proliferation and insulin production
(Matsuda et al., 2018; Tsuji et al., 2014), and have led to the
identification of vitamin D as a potent inducer of cardiomyocyte
proliferation in the context of heart regeneration (Han et al., 2019).
The use of reporter lines to visualize cell cycle, such as the FUCCI
lines (Sugiyama et al., 2009), has proven to be very useful in this
regard as it allows one to assess cell proliferation by fluorescent
protein expression (Han et al., 2019). The establishment of
luciferase promotor lines might also be interesting for screening,
in particular for internal structures (Chen et al., 2013).
Cellular andmolecular mechanisms of organ regeneration in
the zebrafish
So, what has been learned about organ regeneration in the zebrafish
using all these tools? The following section aims to provide an
overview of different sources of progenitor cells that contribute to
rebuild a tissue or organ in this species, and describe some of the
common regulatory mechanisms triggering this process.
Cellular origin
Depending on the tissue, and the degree or nature of damage
involved, newly generated cells can originate from a number of
potential sources (Fig. 3).
Proliferation of existing cell types
In some cases, cells not damaged by the injury can regenerate lost
tissue through proliferation – this may involve a partial de-
differentiation such that the cells enter a proliferative state,
followed by re-differentiation down the same lineage. This is
thought to apply to heart regeneration, where pre-existing
cardiomyocytes de-differentiate and proliferate (Jopling et al.,
2010; Kikuchi et al., 2010) (Fig. 3Ai). Indeed, genetic lineage
tracing using a cardiomyocyte-specific promotor showed that all
newly formed cardiomyocytes at the site of injury are derived from
pre-existent myocardium. Regeneration can also proceed by
proliferation of pre-existing cells, without the need for a
dedifferentiation step before the proliferation phase. This has been
described for liver regeneration – at least when the damage is not too
severe and appears similar to the situation inmammals (Michalopoulos,
2007), where regeneration proceeds via proliferation of pre-existing
hepatocytes (Sadler et al., 2007) (Fig. 3Aii).
Blastema formation
The replacement of a large area of tissue can occur through the
formation of a blastema comprising undifferentiated cells, followed by
cell proliferation and differentiation. A good illustration of this process
is the regeneration of the zebrafish caudal fin. Upon amputation, an
epidermal cap forms that covers the injury. Next, cells from the
amputation plane dedifferentiate forming a blastema under the apical
epidermal cap. During fin regeneration, the dedifferentiated cells
proliferate and undergo re-differentiation to regenerate the various
different cell types that make up the fin (reviewed by Pfefferli and
Jazẃinśka, 2015). The dedifferentiation and proliferation of osteoblasts
close to the amputated area make up the new rays (Stewart and
Stankunas, 2012) (Fig. 3B). However, it has also been shown that fin
regeneration remains viable in the absence of osteoblasts. In this case,
new bone cells can originate from a reserve population of osteoblast
precursor cells (Ando et al., 2017; Singh et al., 2012).
Transdifferentiation
In some cases, regeneration proceeds via dedifferentiation and
proliferation followed by re-differentiation into a different mature
cell type from the starting population. One example of this is the
process following severe hepatocyte loss in the liver. Under these
circumstances, regeneration proceeds initially via the proliferation
of biliary epithelial cells (Fig. 3C), which differentiate into
hepatocytes (Choi et al., 2014). Transdifferentiation has also been
observed in zebrafish pancreatic regeneration. A study searching for
genes that would promote β-cell regeneration identified a protein,
Igfbp1, that potentiated β-cell regeneration by stimulating α- to β-cell
transdifferentiation (Lu et al., 2016).
Differentiation of tissue-resident stem cells/progenitors
Not all organs regenerate through dedifferentiation and proliferation
of mature cells. Injuries to kidney, skeletal muscle or neural tissues
resolve by the proliferation and differentiation of stem cells. For
example, kidney regeneration following chemical damage proceeds
via the proliferation of adult nephron progenitor cells (Diep et al.,
2011). As described in mammals (Wosczyna and Rando, 2018;
Young and John, 2000), skeletal muscle regeneration in zebrafish
requires the proliferation of satellite cells, a population of adult
Pax7+ muscle stem cells that, after an injury to the muscle,
proliferate and fuse with adjacent muscle fibers (Berberoglu et al.,
2017). Regeneration of neurons also has been shown to depend on a
specific progenitor cell population. In the injured zebrafish
telencephalon, new neurons are derived from a subpopulation of
radial glial cells (Kroehne et al., 2011) that have been demonstrated
to function as a stem cells population in uninjured brain tissue
(Rothenaigner et al., 2011) (Fig. 3D).
Environmental cues driving regeneration
Initial wound closure is followed by an inflammatory response
mediated by neutrophils and macrophages, which are attracted by
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signals generated from the dying cells and surrounding tissue, and
home to the site of damage. Indeed, inflammation is essential for organ
regeneration (Mescher, 2017). An inflammatory response precedes
zebrafish brain regeneration (Kyritsis et al., 2012) and has been
suggested to be important for heart regeneration (Lai et al., 2017).
Furthermore, T lymphocytes are involved in the regeneration of several
D
C
i)
ii)
B
A
Fig. 3. Cellular origin of the de novo formed tissue during organ regeneration in the zebrafish. (A) Dedifferentiation, proliferation and re-differentiation.
(Ai) In the heart, cardiomyocytes in close proximity to the injury revert to a less differentiated stage, re-enter the cell cycle and redifferentiate into mature
cardiomyocytes. (Aii) During regeneration of minor liver damage, hepatocyte regeneration occurs with no signs of dedifferentiation prior to cell cycle entry and
proliferation. (B) Blastema formation as an intermediate step during regeneration. After fin amputation, cells of various lineages – including osteoblasts –
dedifferentiate and accumulate under an apical epidermal cap. They then proliferate and redifferentiate to rebuild the missing fin structures. (C) Phenotypic switch
or transdifferentiation during regeneration. Example: after extensive liver damage, biliary ductal cells (green) can transdifferentiate into hepatocytes (green
hexagonal cells) that then differentiate into mature proliferating hepatocytes. (D) Stem cells as progenitor cells. Neural stem cells/progenitor cells proliferate and
differentiate into new neurons during regeneration of the central nervous system. While neuronal regeneration has been well described, less information is
available on robust axon regrowth. Yellow, differentiated cells; orange, dedifferentiated cells; purple, non-osteoblast cells within the fin; green hexagonal cells,
cells undergoing transdifferentiation; blue, stem cells/progenitor cells. Damaged area is shown in gray.
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organs, such as spinal cord, heart and retina (Hui et al., 2017).
Supporting these findings, genetic or chemical ablation of phagocytic
macrophages during regeneration results in impaired fin (Li et al.,
2012; Petrie et al., 2014), lateral line (Carrillo et al., 2016), heart
(Lai et al., 2017) and spinal cord (Tsarouchas et al., 2018) regeneration.
The ECM and the stromal environment also influence regeneration.
Not only do they provide structural support for progenitor cells, but
they also have an impact on their proliferation and differentiation.
Indeed, fibrosis and regeneration are tightly linked processes. For
example, the pro-fibrotic molecule TGFβ induces not only fibrosis but
also cardiomyocyte proliferation (Chablais and Jazwinska, 2012).
Moreover, ablation of collagen-producing cells impairs cardiomyocyte
proliferation after heart injury (Sánchez-Iranzo et al., 2018), and ECM
proteins are required for spinal cord regeneration (Mokalled et al.,
2016; Wehner et al., 2017).
The nervous system is also key for adequate regeneration. This
has been demonstrated both in the fin, where denervation following
amputation results in impaired regeneration (Simões et al., 2014),
and in the heart, where regeneration is also impaired following
inhibition of peripheral nervous system activity (Mahmoud et al.,
2015). Future work is needed to further unravel the role of the
nervous system during organ regeneration, which can range from
secretion of molecules to establishment of synaptic connections.
Finally, revascularization is also essential for proper regeneration.
When this process is impaired during cardiac injury, regeneration is
blocked. Studies conducted on this subject suggest that revascularization
of the injured area is essential to stimulate cardiomyocyte
proliferation (Harrison et al., 2015; Marín-Juez et al., 2016). The
exact mechanisms through which revascularization contributes to
regeneration remain to be discovered.
Molecular pathways promoting regeneration
The key signaling pathways involved in embryonic development are
often also involved in organ regeneration. To study their function,
researchers have used transgenic lines for temporal and tissue-
specific overexpression of a core gene of a particular pathway and
studied the effect on organ regeneration. In addition, analysis of
organ regeneration in mutants of a gene involved in the pathway
of interest has been informative. Here, we provide a few examples of
how such pathways contribute to regenerative processes in
zebrafish; we refer the reader to the papers cited for further
details. One example of a pathway important for regeneration is the
retinoic acid (RA) pathway. RA regulates FGF, Wnt/β-catenin and
Igf signaling during blastema formation in the regenerating caudal
fin (Blum and Begemann, 2012), and stimulates cardiomyocyte
proliferation, likely in a paracrine manner (Kikuchi et al., 2011).
The fibroblast growth factor (FGF) and bone morphogenetic protein
(BMP) pathways, which are essential for organ growth and
differentiation (Kan et al., 2009; Smith et al., 2006), are also
involved in regeneration. BMP signaling has been implicated in the
regeneration of several organs, including the liver, the heart and the
fin (Kan et al., 2009; Sehring et al., 2016). Similarly, FGF is
involved in multiple processes of regeneration, including the control
of blastema proliferation during fin regeneration (Lee et al., 2005;
Poss et al., 2000) or heart regeneration (Lepilina et al., 2006). One of
the most universally used pathways in tissue regeneration
throughout the animal kingdom is the Wnt signaling pathway
(Slack, 2017). In the zebrafish, it regulates blastema proliferation in
amputated caudal fin among many other processes (Stoick-Cooper
et al., 2007; Wehner et al., 2017). Juxtacrine signaling pathways,
such as the Notch pathway are also associated with proliferation
control both in fin (Münch et al., 2013) and heart (Zhao et al., 2014).
Other pathways involved in the control of cell cycle include the
Hippo-Yap/TAZ pathway (Flinn et al., 2018; Mateus et al., 2015),
which has recently received considerable attention in the field of
regeneration because it links the effect of a changing environment
and mechanical forces in response to injury with the regenerative
response in the surrounding cells. In all cases, specificity and temporal
control of these pathways is crucial for promoting regeneration. In
addition, depending on the combination of active pathways, different
cellular responses are elicited, ranging from cell differentiation or
proliferation to migration to epithelial-to mesenchymal transition, all
of which are crucial for organ regeneration.
In sum, during zebrafish regeneration, cells that repopulate
injured areas can have different origins, depending not only on the
lesioned organ, but also on the extent and the type of injury.
Furthermore, for regeneration to succeed there must be a balance
between the signals that induce cell proliferation and those that halt
cell division and determine cell fate.
Extent and limits of the regenerative capacity in zebrafish
Without doubt, the zebrafish is an organism with an amazing
capacity for regeneration. Here, we discuss some of the mechanisms
that might underlie this capacity, but also highlight some possible
limitations of organ regeneration in this animal.
Possible explanations for the high regenerative capacity
The regenerative capacity in the zebrafish might rely on its capacity to
activate specific gene regulatory networks in response to injury that
are repressed in other animal models (Yang and Kang, 2018). Indeed,
epigenetic modifications are necessary to trigger regeneration. One of
the earliest findings in this field reported that, in the fin, histone
modifications precede initiation of regeneration (Stewart et al., 2009),
and histone methyltransferases are necessary for this process (Dupret
et al., 2017). The use of ChIPseq and transgenesis has provided much
insight into the landscape of active enhancers during organ
regeneration, identifying enhancer elements that are activated in
response to injury consistently in different organs and even in different
organisms (Kang et al., 2016; Pfefferli and Jazẃinśka, 2017).
One important characteristic of zebrafish is its capacity to grow
throughout its lifespan, meaning that most of the zebrafish cells, e.g.
cardiomyocytes, retain their proliferative capacity (Wills et al.,
2008). For heart regeneration, cardiomyocyte ploidy has been
shown to be a limiting step. Zebrafish cardiomyocytes are
mononuclear and diploid, whereas in adult humans they are
mostly polyploid. Indeed, induction of polyploidy in zebrafish
cardiomyocytes abrogates heart regeneration (González-Rosa et al.,
2018; Patterson et al., 2017).
The immune system might play a key role in defining the
regenerative capacity of an organism. As an example, differences in
immune response to cardiac injury in two teleost models (medaka
and zebrafish) correlate with reduced regenerative capacity of
medaka (Lai et al., 2017). Finally, regenerative capacities might also
be attributed to differences in the environment that species inhabit –
exemplified by the differences in heart regeneration in surface
versus cave morphs of the cavefish Astyanax mexicanus (Stockdale
et al., 2018). Thus, using other fish models with different tissue
recovery capacities for regeneration studies (see Table 4) may
contribute to a better understanding of the different mechanisms
underlying this process.
Unlimited regeneration in the zebrafish?
Regeneration in the zebrafish seems to be nearly unlimited:
regeneration of the caudal fin and barbell occurs even after
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repetitive amputations (Azevedo et al., 2011; LeClair and
Topczewski, 2010), although subtle changes to the newly formed
organ, such as variation in the pigmentation patterning of the fin or
the position of the bony ray bifurcations, have been observed
(Azevedo et al., 2012). Moreover, age does not seem to limit
regenerative capacity, at least for the heart (Itou et al., 2012).
Nevertheless, there is evidence that organ regeneration can be
partially incomplete. For example, although heart morphology is
restored after lesion and no scar is visible, the ventricular wall
motion of the regenerated area remains partially affected and
contracts asynchronously with the remainder of the heart, even after
complete tissue regeneration (González-Rosa et al., 2014).
Furthermore, following resorption of extracellular fibrotic
components during heart healing, accumulated fibroblasts are not
fully eliminated after complete regeneration but become inactivated
(Sánchez-Iranzo et al., 2018). It is thus important to distinguish
between morphological and functional regeneration and to
recognize that – even in a system such as zebrafish – regeneration
may not be functionally ‘perfect’.
Conclusions and perspectives
In conclusion, using the zebrafish as an animal model in regeneration
studies has undoubtedly contributed to the development of the field
but, as with any other model, there are advantages and disadvantages
to consider (Table 5). Despite the contributions made, there are still
unanswered questions. For example, there is little information on how
metabolism is linked to regenerative capacity. Furthermore, the
interplay between fibrosis and regeneration, as well as the
mechanisms of fibrotic tissue regression, are still not fully
understood. Why is a scar formed under some circumstances or in
some organs while in other cases regeneration occurs? Towhat extent
does regeneration rely on a transient fibrotic scar? It is also important
to understand howmechanical forces influence regeneration – studies
in the zebrafish might help to explain how, for example, stiffness or
shear stress regulates organ regeneration. In addition, it is intriguing
that regenerating tissues stop growing at the appropriate size; this is
poorly understood. Indeed, there is still a dearth of knowledge
regarding which cell populations become active during regeneration
and whether all the cells have the same regenerative potential. To
answer many of these questions, some of the molecular tools such as
conditional genome editing will need to be improved and widely
established for regeneration studies. Advances in intra vital imaging
for adult zebrafish will also help to expand our toolbox. With these
new technological advances in hand, important discoveries on the
mechanisms of organ regeneration using the zebrafish model will
certainly be achieved. A deep knowledge on the mechanisms of
organ regeneration in species such as the zebrafish with innate
regenerative capacity might serve as inspiration for the development
of therapeutic strategies in mammals.
Acknowledgements
We thank Andres Sanz for critical reading of the manuscript and the reviewers for
their suggestions. We also acknowledge all those authors whose work could not be
cited here due to space limitations. Their research keeps providing us with invaluable
knowledge in the field of zebrafish regeneration.
Competing interests
The authors declare no competing or financial interests.
Funding
N.M. is supported by a European Research Council starting grant (2013 zebraHeart-
337703), E.L. is supported by the European Commission (H2020-MSCA-ITN-2016
European Industrial Doctorate 4DHeart 722427).
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Table 5. Advantages and disadvantages of the zebrafish as a
regeneration model
Advantages Disadvantages
High regenerative capacity
Vertebrate model
Established husbandry
protocols
Low-cost maintenance
External development
Possibility of working with large
numbers
Ease of in vivo imaging during
development
Genetic engineering, annotated
genome
Physiological similarities with
humans (e.g. heart rate)
Non-mammalian model
Partial genome duplication
Dearth of antibodies for protein studies
Difficulty in performing in vivo imaging
in adult stages
Difficulty in using some molecular
tools (e.g. knock-ins)
Anatomical and physiological differences
to humans (e.g. two-chambered heart)
Table 4. Other teleost fish models of regeneration
Fish model
Organs in which regeneration has been
studied
Astyanax mexicanus (Mexican
cave fish)
Heart (Stockdale et al., 2018)
Carassius auratus (Goldfish) Spinal cord (Bernstein and Gelderd,
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Eye (Parrilla et al., 2012)
Fin (Caskey and O’Brien, 1948)
Heart (Grivas et al., 2014)
Oryzias latipes (Medaka) Kidney (Watanabe et al., 2009)
Fin (Nemoto et al., 2007)
Pancreatic β-cells (Otsuka and Takeda,
2017)
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