As one of the most vital organs, mammalian hearts have a minimal capacity for regeneration upon disease or injury in the post-natal period, especially for replenishing cardiac muscle cells (cardiomyocytes, CMs). Instead, the infarcted hearts undergo fibrotic repair, which in turn deteriorates tissue contractility and function, eventually leading to heart failure and organismal death (Kong et al., 2014). In contrast to the limited regenerative capacity of adult mammals, certain fishes and amphibians, and even neonatal mice can regenerate their hearts after injury (Vivien et al., 2016). This is an excellent example of how the regenerative capacity of hearts exists unevenly across species and developmental stages, as mammals possess the regenerative capacity only for a short time window after birth (Porrello et al., 2011; Haubner et al., 2012, 2016). These mammalian and non-mammalian models provide unique opportunities to study the intrinsic capacity and mechanisms of heart regeneration. Amongst, extensive knowledge was gained from the zebrafish studies. Since the ground-breaking discovery of zebrafish heart regeneration by Poss et al. (2002), researchers have made in-depth investigations to understand the mechanisms of heart regeneration in zebrafish (Marques et al., 2019; Jaźwińska and Blanchoud, 2020; Potts et al., 2021). Briefly, zebrafish hearts mount a robust immune response in the recruitment of macrophages and neutrophils immediately after injury (Lai et al., 2017) and fast revascularization that expands superficially and intraventricularly and serve as the scaffold for CM repopulation (Marín-Juez et al., 2016, 2019). Almost concurrently, epicardium activates and expands by proliferation to cover the injured area and serve as a signaling hub to stimulate CM de-differentiation and proliferation in the border zone of the injured area (Kikuchi et al., 2010, 2011a,b; Jopling et al., 2011; González-Rosa et al., 2012; Cao and Poss, 2018). Newly formed CMs gradually replace scar tissue coincident with ECM remodeling (Sanchez-Iranzo et al., 2018) and scar resolution (Bevan et al., 2020; Simoes et al., 2020), eventually restoring the morphology and function of the heart. Conserved processes have been shown in neonatal mouse heart regeneration, particularly the source of regenerated CMs (Porrello et al., 2011; Vivien et al., 2016). Furthermore, hints gained from zebrafish studies have been applied to mice models to accelerate cardiac repair (Chen et al., 2016; Honkoop et al., 2019). A comparative study in zebrafish and mouse injured hearts even revealed microRNA dynamics that may regulate CM proliferation and cardiac repair (Crippa et al., 2016). However, considering the taxonomy distance between zebrafish and mice, a comparative approach may be more feasible for more closely related species with similar physiology and structure.

Unlike zebrafish, medaka showed impaired heart regeneration, indicated by a lack of revascularization, low CM proliferation, and a permanent fibrotic scar in the injured area after resection (Ito et al., 2014). To understand the differences in cardiac repair in zebrafish and medaka, Lai and colleagues performed a global transcriptomic analysis and revealed a robust immune response and angiogenic revascularization exist preferentially in zebrafish (Lai et al., 2017). Coincidently, they observed a reduced macrophage infiltration and prolonged neutrophil recruitment/retention in medaka hearts compared to zebrafish. The blunted immune response in medaka encouraged the investigation of the acute immune response and timely macrophage recruitment in heart regeneration. Indeed, in a loss-of-function setting, delayed macrophage recruitment by clodronate liposome pre-depletion abolished the regenerative capacity in zebrafish (Lai et al., 2017), which correspond nicely with findings in neonatal mice (Aurora et al., 2014). These results support an essential role of macrophage function in heart regeneration across species and encore the requirement of the macrophage function in the regeneration of other organs, including fin, retina, optic tectum, brain, and spinal cord reviewed elsewhere (Var and Byrd-Jacobs, 2020). They further identified the immunostimulant poly I:C as one of the upstream candidates that may trigger the differential transcriptomic response found between zebrafish and medaka. Indeed, they further showed that stimulating immune response by poly I:C administration promotes heart regeneration in medaka in a gain-of-function setting (Lai et al., 2017). These results support the strength of comparative analysis using fish models to gain knowledge conserved across species and identify pro-regenerative factors. However, it remains unclear how poly I:C (or other immunostimulants) promote heart regeneration in medaka and whether the same principle may apply to mammals, awaiting further investigation.

The capacity to regenerate retinal neurons after injury also varies drastically among vertebrate species. While mammalian Müller glia (MG) do not spontaneously regenerate lost retinal neurons, zebrafish MG cells possess a robust capacity to regenerate all retinal cell types and recover their visual ability (Vihtelic and Hyde, 2000; Sherpa et al., 2008; Goldman, 2014; Gorsuch and Hyde, 2014; Lenkowski and Raymond, 2014). Several pluripotent factors, including ascl1a, lin-28, and sox2, regulate the dedifferentiation, reprogramming, and proliferation of MG cells into various retinal cell types during retina regeneration in zebrafish (Ramachandran et al., 2010a; Gorsuch et al., 2017). Sox2 is also one of the four Yamanaka factors that induced pluripotent stem cell status (Takahashi et al., 2007). In addition to retina, retinal pigment epithelium (RPE) regeneration was also recently described in zebrafish (Leach et al., 2021). Similar to heart regeneration, the immune response, particularly the macrophages and microglia cells, responds to injury and plays a critical role in retina and RPE regeneration, potentially associated with phagocytotic debris clearance and cytokine secretion (Mitchell et al., 2019; Leach et al., 2021).

During retina development, medaka neural stem cells behave similarly to those in zebrafish (Martinez-Morales et al., 2009; Centanin et al., 2011, 2014). However, Lust and Wittbrodt discovered that medaka showed limited regenerative capacity in the retina. The MG cells proliferate but fail to self-renew and reprogram, eventually giving rise to only photoreceptor cells (Lust and Wittbrodt, 2018). Moreover, by comparing medaka with zebrafish, they identified that medaka MG cells fail to maintain sox2 expression after injury and demonstrated that sustained sox2 expression in medaka MGs confers regenerative response (Lust and Wittbrodt, 2018). Similar to the above-mentioned reciprocal analyses in heart regeneration, Lust and Wittbrodt were able to identify the critical factor and demonstrated the functional relevance of sox2 expression in promoting retina regeneration.

More recently, Hoang et al. (2020) identified the evolutionarily conserved and species-specific gene regulatory networks that control the quiescent, reactive, and proliferative MG transition after retinal injury in another cross-species comparison between mice, chick, and zebrafish. They further demonstrated that deleting the factors maintaining the quiescent state may promote MG reprogramming into regeneration-competent cells in adult mice (Hoang et al., 2020).

Unlike mammals, zebrafish respond to injury or degeneration by inducing specific neurogenic programs and constitutive neurogenesis for tissue regeneration (Diotel et al., 2020). Learning the regenerative mechanisms occurring in zebrafish will be invaluable for developing therapeutics for brain injury and degenerative diseases. In contrast to target-oriented studies, new knowledge may come from side-by-side and unbiased comparisons of animal models with divergent regenerative capacities. Unfortunately, mice brains and zebrafish brains show distinct features other than regenerative capacity, including the overall anatomy and neurogenic niches, thus preventing direct comparisons (Diotel et al., 2020; Labusch et al., 2020). Therefore, it is relevant to explore the mechanisms of brain regeneration in more closely related models.

The regenerative capacity of zebrafish central nervous system (CNS) has been investigated in the optic tectum (Ito et al., 2010; Shimizu et al., 2018; Lindsey et al., 2019) and telencephalon (Kroehne et al., 2011; Marz et al., 2011; Kishimoto et al., 2012). In the adult zebrafish CNS, both MG and the radial glia (RG) cells are activated to proliferate and differentiate into new neuronal cells following injury (Raymond et al., 2006; Ito et al., 2010; Shimizu et al., 2018). These regenerative responses seem to be induced and facilitated by immune responses (Kyritsis et al., 2012; Caldwell et al., 2019), while a specific inflammatory signaling cascade is stimulated by microglia during zebrafish brain repair (Kanagaraj et al., 2020).

A comparative study of brain regeneration in medaka and zebrafish has also been reported very recently (Shimizu and Kawasaki, 2021). Medaka shares a similar brain structure with zebrafish and neural stem cells (NSCs) niche for brain development and growth (Adolf et al., 2006; Grandel et al., 2006; Alunni et al., 2010; Kuroyanagi et al., 2010). Main NSCs exist in the optic tectum of both zebrafish and medaka, including the proliferative neuroepithelial-like stem (NE) cells and the quiescent RG cells (Alunni et al., 2010; Ito et al., 2010; Takeuchi and Okubo, 2013; Dambroise et al., 2017). However, medaka could not regenerate their optic tectum after stab injury and thus leaving a permanent scar (Shimizu and Kawasaki, 2021). In medaka, RG cells were similarly activated for proliferation upon tectum injury, but they failed to differentiate into neuron cells. Unlike the scenario in the retina, sox2 is substantially expressed in both zebrafish and medaka optic tectum and does not associate with the differential regenerative ability. Instead, the expression of pro-regenerative transcriptional factors ascl1a and oct4 were missing in the medaka. As a result, glial scar-like structures composed of GFAP+ radial fibers filled the injured area of the medaka optical tectum. Follow-up studies might be required to test the functional relevance of the ectopic expression of ascl1a and oct4 in promoting RG differentiation and optic tectum regeneration in medaka.

Among various tissues and organs, appendage regeneration draws major attention early on in the field as teleost fish, urodeles, and amphibians all can regenerate their fins, arms, and legs following amputation (Daponte et al., 2021). Compared to limbs of urodeles and amphibians, fish fin structure is simpler and consists of bony fin rays covered by thin epidermal cells (Grandel and Schulte-Merker, 1998). Nevertheless, limb and fin are homologous tissue across vertebrate species (Yano and Tamura, 2013). A forward-genetic screen done in zebrafish identified a novel and conserved regulator of appendage patterning. When mutated, zebrafish formed limb-like bones in fins, suggesting the conservation in skeleton development and the potential of fin-to-limb transition (Hawkins et al., 2021). The regenerative capacity of the fish fin was examined as early as the 1700s to understand appendage regeneration with the first reports by French naturalist Broussonet (Broussonet, 1786; Broussonet, 1789). Like the limb regeneration in urodeles and amphibians, zebrafish repair their caudal fin by blastema-mediated epimorphic regeneration (Poss et al., 2003). In this context, blastema originates from dedifferentiated mesenchymal cells and is the primary source for growing new tissues, including bone, nerve, and vessel (Poss et al., 2003; Straube and Tanaka, 2006; Pfefferli and Jazwinska, 2015). Conserved pathways, including the Wnt/β-catenin pathway, were shown to regulate appendage regeneration across different vertebrate species, including zebrafish, Xenopus, and axolotl (Kawakami et al., 2006; Yokoyama et al., 2007).

Like zebrafish, medaka also regenerates their fin after amputation via blastema-mediated epimorphic regeneration (Katogi et al., 2004; Nakatani et al., 2007). It will be interesting to learn if medaka fin regeneration shares a conserved regenerative program with zebrafish in a cross-species study. In addition, inflammation and macrophages play a central role in both heart and fin regeneration in zebrafish but seem deficient/blunt in medaka hearts upon injury (Petrie et al., 2014; Lai et al., 2017). One may wonder whether systemic inflammation and immune response contribute differently upon fin and heart injury in medaka, which will be further discussed and await future investigation.

Overall, all these studies highlight the strength of inter-species comparisons between zebrafish (regenerative) and medaka (non-regenerative) organs to identify (and in some cases also to validate) the potential triggers of tissue regeneration. It is worth mentioning that other comparisons of organisms/conditions exhibiting diverse regenerative capacity have also been reported. For example, Stockdale and colleagues identified genes fundamental to heart regeneration by comparing the injury response of regenerative Astyanax mexicanus surface fish with their non-regenerative counterparts Pachón cave-dwelling fish (Stockdale et al., 2018). Following such regenerative traits among other species, researchers have also explored the evolutionary concept of regeneration amongst other teleost species (Table 1). In addition, comparing the transcriptomes and open chromatin landscapes of the cardiac cells isolated from the regenerative neonatal vs. non-regenerative adult mice hearts, Wang Z. et al. (2020) revealed the gene regulatory networks in diverse cardiac cell types and extracellular mediators for cardiomyocyte proliferation, angiogenesis, and fibroblast activation. Furthermore, mammals like the African spiny mouse (Acomys) can regenerate their ear, skin, heart, and bones in contrast to the house mouse (Mus musculus), providing more opportunities for cross-species analyses (Seifert et al., 2012; Matias Santos et al., 2016; Simkin et al., 2017; Qi et al., 2021).

Comparing organisms with similar regenerative properties may also identify the conserved regenerative programs. For example, from an inter-species comparison of zebrafish and African killifish (Nothobranchius furzeri) following fin amputation, Wang and colleagues identified the evolutionary conserved regenerative response elements (RRE) (Wang W. et al., 2020). Activation of inhba, a gene downstream of the RRE, is essential for both fin and heart regeneration and requires the binding motifs of activator protein 1 (AP-1) complex. Such enhancer is also present in mammals, shares Ap-1 binding motifs, and responds to injury, although it cannot promote regeneration. These results suggest that RREs might have been repurposed in regeneration-incompetent animals during evolution and only promote tissue repair but not regeneration (Yang and Kang, 2019; Wang W. et al., 2020). Of note, killifish possess the regenerative capacity in multiple organs, including the heart (Wang W. et al., 2020), fin (Wendler et al., 2015), and brain (Van Houcke et al., 2021) while being phylogenetically closer to medaka (Terzibasi et al., 2007), represent an alternative model for inter-species comparison.

In addition to the availability of progenitor/stem cell populations in each tissue/organ, we wonder how systemic responses, including immune response, neural innervation, hormonal regulation, metabolic shift, contribute differently to the respective injured tissues/organs of the same organism and lead to uneven regenerative capacity. The problem is especially apparent when one teleost zebrafish can regenerate organs such as the heart, retina, and brain, while another teleost medaka cannot. Taking the heart for example, the blunt immune response seems to be the major obstacle for medaka to initiate regenerative programs, but how does the same systemic (immune) response sustain the regeneration of other organs, for example, the fin? Are there tissue-specific contributions/responses, for example, residential immune cells, tissue-specific injury response elements, or even changes in the epigenomic landscape? These questions may be best addressed in medaka where the organ-specific regenerative capacities are uneven and well studied, including the regenerative fin (Katogi et al., 2004), kidney (Watanabe et al., 2009), liver (Van Wettere et al., 2013), and pancreas (Otsuka and Takeda, 2017), and non-regenerative heart (Ito et al., 2014; Lai et al., 2017), retina (Lust and Wittbrodt, 2018), and brain (Shimizu and Kawasaki, 2021). The potential findings can be cross-species compared and further validated in zebrafish loss-of-function and medaka gain-of-function experiments. A similar concept could also apply to other species. Differential regenerative capacity can be observed in lower vertebrates where some lizards can replace their tail but not their limb (Alibardi and Toni, 2005). Even in mice, the uneven regenerative capacity exists ranging from active (intestine and skin), partial (liver, pancreas, muscle), to none (CNS, heart, and most other internal organs) (Iismaa et al., 2018; Figure 1).

Even between two regenerative organs, researchers have identified common and tissue-specific regeneration responsive elements/enhancers by inter-organ comparisons. For example, Kang et al. (2016) have performed such a comparative study identifying a tissue regeneration enhancer element (TREE) from zebrafish heart and fin, which locates upstream of lepb gene and activates following injury. This element could also be activated in neonatal mouse tissues upon injury and may be engineered to modulate the regenerative potential of vertebrate organs. Another study by Pfefferli and Jaźwińska (2017) identified a 3.18 kb regulatory element upstream of ctgfa gene, named as careg element that drives the regenerative response in both zebrafish fin and heart via TGFβ/Activin-β signaling pathway. This type of study paved the way for identifying evolutionarily conserved RREs, which can also be analyzed in medaka to decipher how these RREs exist and regulate the regenerative programs in a tissue-specific manner.

Various injury models have been established in fish models to introduce tissue injury and investigate the reparative process. In addition to the feasibility and reproducibility, these methods were often established based on the similarity to the human diseases or trauma conditions to gain translational value (Table 2). The resection or amputation model is one of the most commonly used injury models, involving surgical removal of a part of the tissue for observing the restoration of size, morphology/structure, and function of the injured tissue. Resection is straightforward, cheap, and reproducible and has been widely adopted to fin (Poss et al., 2003; Nakatani et al., 2007) and heart (Poss et al., 2002; Ito et al., 2014) regeneration studies in both zebrafish and medaka. However, resection is often accompanied by excessive bleeding and open wounds, which leads to infection and high mortality. Also, resection is sometimes infeasible for internal organs or tissues those unexposed or too small. Similar models include stabbing and transection, which works by surgically disrupting the integrity without removing any tissue and are well established in the retina, brain, and spinal cord regeneration in both zebrafish (Becker et al., 1997; Marz et al., 2011; Shimizu et al., 2018) and medaka (Shimizu and Kawasaki, 2021).

Slightly different from resection, researchers adapted the cauterization method in fish models to mimic the fibrotic repair in mammalian organs, which is highly associated with the inflammatory response (Strungs et al., 2013; Polizzotti et al., 2016). The cauterization technique involves burning or freezing the target tissue with electric or metal probes and introducing necrotic and apoptotic cell death. Cryoinjury is popular in heart regeneration studies in both zebrafish (Gonzalez-Rosa et al., 2011; Schnabel et al., 2011; Dyck et al., 2020) and medaka (Lai et al., 2017) as it mimics the myocardial infarction in mammals better than resection model (Chablais et al., 2011; Darehzereshki et al., 2015). This technique can also be applied to external organs, such as the fin (Chassot et al., 2016) and gills (Ramel et al., 2021). However, cauterization is technically challenging and less reproducible compared to resection. Also, the wound usually takes a longer time to recover as the cell debris needs to be cleared before regeneration occurs (Schnabel et al., 2011). To increase the accuracy and reproducibility, cauterization can also be performed by using high-powered lasers in fish tissues, including the retina (Conedera et al., 2017; Lust and Wittbrodt, 2018) and skin (Richardson et al., 2013). The target tissue has to be exposed or transparent for laser penetration in this case. Overall, most physical injuries are invasive to cause high mortality and take practice to be consistent.

Apart from physical injury models, genetic cell ablation models in fish were established by expressing enzymes that catabolize cytotoxic products added to the system or induce cell death directly (Table 2). The former method was developed by expressing the bacterial enzyme Nitroreductase (NTR), which alone is not toxic but can catabolize the prodrug metronidazole (Mtz) to induce cytotoxicity (Lindmark and Müller, 1976). This system can achieve spatial (tissue-specific expression of NTR) and temporal control (the timing of adding Mtz), and labeling the target cells (co-expression with reporter system) at the same time. NTR/Mtz system has been used to tease out the functions of specific cell types in a complex process of organ regeneration in zebrafish, including the heart (Curado et al., 2007; Wang et al., 2013; Zhang et al., 2013), fin (Petrie et al., 2014), pancreatic β-cells (Pisharath et al., 2007), bone (Willems et al., 2012) and RPE regeneration (Hanovice et al., 2019). This system is also applicable to medaka, demonstrated by accessing regeneration capacity using NTR/Mtz mediated genetic ablation of the pancreatic β-cell population (Otsuka and Takeda, 2017), osteoblasts (Willems et al., 2012), and bone progenitor cells (Dasyani et al., 2019) in fin regeneration.

The latter genetic ablation tool to study zebrafish development and regeneration involves diphtheria toxin A (DTA) expression under a tissue-specific promoter, exampled by crystallin promoter-driven DTA expression in lens (Kurita et al., 2003), elastase A promoter-driven DTA expression in exocrine pancreas (Wan et al., 2006), and myl7 promoter-driven DTA expression in cardiomyocytes (Wang J. et al., 2011). Though the DTA approach lacks temporal control for activation, it is highly toxic for killing the target cells efficiently. In a modified method, the temporal control can be achieved by expressing the human diphtheria toxin receptor (DTR) and further activate cytotoxicity by diphtheria toxin injection (Jimenez et al., 2021). The genetic ablation models are technically simpler with faster recovery. These models can be used combined with physical injury to tease the role of a specific cell type during complex organ regeneration. For example, Sanchez-Iranzo et al. (2018) depleted fibroblast cells after cardiac cryoinjury and demonstrated how fibroblasts contribute to heart regeneration. Overall, most of these injury models developed in zebrafish may also apply to medaka in comparative studies of tissue regeneration.

One of the best attributes of using zebrafish and medaka as research models is the tools and techniques available for visualizing specific cell types and biological processes in vivo. Here, we summarize the visualization tools that have been established in fish models, especially in zebrafish and correspondingly in medaka, with a focus on the transgenic reporter lines (Table 3) and alternative approches.

Since fluorescence reporter driven by tissue-specific gene promoter is applicable and efficient in zebrafish and medaka, many transgenic lines have been generated to study specific tissues/organs in development and diseases. Here we summarize tissue-specific reporters described in major tissue regeneration studies previously mentioned in Table 3. In addition to tissue-specific reporter lines, researchers can also use antibodies against cell-specific transcription factors or cytosolic proteins to assess cellular dynamics in growth, development, and regeneration. For example, Mef-2 (sc-313), nkx2.5 (GTX128357) or MF20 (Fischman, D.A., DSHB) antibody can be used in combination with proliferation/cell cycle markers PCNA (GTX124496), Anti-phospho-Histone H3 (Ser10) (06-670, Merck), and BrdU/EdU (C10086, ThermoFisher, Eugene, OR, United States) to label the proliferating CMs (Chablais et al., 2011; Chablais and Jaźwińska, 2012a). Similarly, transgenic medaka lines have also been generated to facilitate cardiac research using zebrafish cmlc2 regulatory elements for myocardial expression (Taneda et al., 2010). In addition to labeling cardiac tissues, researchers have successfully developed medaka reporters utilizing zebrafish skeletal specific mylz2 promoter to label skeletal muscles (Zeng et al., 2005). Vice versa, medaka mylz2 promoter can also recapitulate GFP expression in zebrafish (Zeng et al., 2005). On the same note, medaka β-actin promoter can drive ubiquitous gene expression in both medaka and zebrafish (Yoshinari et al., 2012), while zebrafish krt8 promoter can label both skin and intestinal epithelium in medaka, as almost identical to zebrafish (Zeng et al., 2005). These examples showed that the transcriptional regulation of many genes is highly conserved in both species, with many tissue-specific transgenes developed to facilitate research, as summarized in Table 3.

Restoring vasculature and circulation is one of the first steps during tissue regeneration (Jung and Kleinheinz, 2013). Both these blood vessels and lymphatic vessels that regulate tissue homeostasis and immune cell trafficking can be visualized by reporters and have been used extensively for regenerative studies in zebrafish (Table 3). To name a few of the most commonly used reporter lines, tie2 reporter for pan-endothelial cells (Motoike et al., 2000), fli1a reporter for endothelial and endocardial cells (Lawson and Weinstein, 2002), kdrl reporter for arterial vessels (Wang et al., 2010), and lyve1 (Okuda et al., 2012) reporter for lymphatic vessels in zebrafish. Correspondingly, a wide range of transgenic reporter strains has been generated in medaka based on orthologous gene promoters (Table 3). Moreover, one can also use staining methods to label the vasculature when transgenic animals are inaccessible. One of the widely adopted vasculature staining methods involves alkaline phosphatase (AP) staining, which relies on the endogenous AP activity to convert NBT/BCIP into purple precipitates in endothelial cells for rapid visualization in larvae (Childs et al., 2002) and heart (Lai et al., 2017). In addition, the Fli1 antibody (ab133485) can be used to mark endothelial cell nuclei in zebrafish by immunostaining (Bensimon-Brito et al., 2020). Aside from post-fixation staining, angiography can be achieved by fluorescent dextran injection into the circulation of larval (Hoeppner et al., 2015; Takanezawa et al., 2021) and adult fish (Pugach et al., 2009).

Immune response, especially inflammatory cell infiltration and resolution, is a critical component of tissue regeneration to prevent infection, clear damaged tissue, maintain tissue integrity, and sometimes even is associated with the fibrotic response and cell proliferation (Julier et al., 2017). On top of the tremendous capacity in regeneration, zebrafish possess both innate and adaptive immunity comparable to mammals (Trede et al., 2004), making it a powerful model to study the role of immune response in tissue repair and regeneration (Var and Byrd-Jacobs, 2020). Inflammatory cells, including neutrophils and macrophages, are among the first responders recruited to the injured tissue by chemokines and damage-associated molecular patterns (DAMPs) (McDonald et al., 2010; Soehnlein and Lindbom, 2010). Some of the zebrafish reporter lines have been used to visualize these innate immune cells, including fluorescent genes expression driven by neutrophil-specific mpx promoter (Renshaw et al., 2006) and macrophages specific mpeg1.1/mpeg1.4 (Ellett et al., 2011) and mfap4 (Walton et al., 2015) promoters. Taking advantage of the transparent tissue at the larval stage, zebrafish have been extensively used for studying the dynamic and function of these inflammatory cells in tissue repair/regeneration (Li et al., 2012). Apart from the innate immune system, some adaptive immune cell reporters were also established in zebrafish (Table 3). Materials for zebrafish immune research have also been previously reviewed (Martins et al., 2019). The immune system in medaka is less studied compared to zebrafish. Still, some immune cell reporters have been generated in medaka based on zebrafish orthologous genes (Table 3), including neutrophil-specific mpo reporter (alias to mpx) (Grabher et al., 2007; Crespo et al., 2014), macrophages specific mpeg1.1 reporter (Phan et al., 2020), and pan mononuclear phagocytes cxcr3.2 reporter (Aghaallaei et al., 2010). For the adaptive immune cells, medaka gained interest for studying T-cell development where Tg lines were developed respectively (Bajoghli et al., 2019). Given the importance of immune response in tissue repair/regeneration and the amount of knowledge gained in zebrafish, it is pretty evident that corresponding transgenic reporter lines in medaka await future development for comparative studies. Specifically, it would be interesting to learn more about the critical roles of immune cells that plays similarly or differently in these two model systems.

Due to the limited resource of antibodies against fish proteins and reporters for labeling immune cells, other approaches can be applied to label and even isolate immune cells. For example, isolectin B4 (IB4) and liposome-uptake may label macrophage and other phagocytes in both zebrafish and medaka. In a comparative study, IB4 labels mainly macrophages in zebrafish and medaka and show colocalized signals with zebrafish mpeg1 reporter signals (Lai et al., 2017). On the other hand, DiI liposomes can label the phagocytes efficiently in both zebrafish and medaka based on their properties of macrophage ablation when loaded with clodronate (Lai et al., 2017). Despite in limited numbers, some antibodies work in both fish models in labeling the immune cells, including Lcp1 (GTX124420) (Redd et al., 2006), Lyz (GTX132379) for leukocytes, Spi/Pu.1 for myeloid cells (GTX128266), Mpx (GTX128379) for neutrophils (Lai et al., 2017), Mpeg1 (GTX54246) for macrophages (Simoes et al., 2020), and anti-4C4 for microglia (Becker and Becker, 2001).

Here, we have tabulated the most widely used fish reporter and transgenic lines in Table 3. More transgenic lines that label different tissues can be easily looked up in the Zebrafish Information Network (ZFIN¹) and accessed from the Zebrafish International Resource Center (ZIRC²) and the European Zebrafish Resource Center (EZRC³), or the NBRP Medaka website⁴. Overall, common regulatory elements between zebrafish and medaka support that the activation of zebrafish-specific factors can be well recapitulated in the medaka and vice-versa. This evidence further highlights the conserved gene-regulatory networks between zebrafish and medaka, making them excellent models to perform comparative studies in tissue regeneration.

Taking advantage of being model organisms and a broad research community, zebrafish and medaka are well-equipped with tools for genetic manipulations for generating animal models for specific cell ablations, visualization, and functional manipulations introduced previously. Here, we summarize these toolsets established in zebrafish and medaka (Table 4) to investigate the cellular and molecular mechanisms of tissue regeneration.

Genetic manipulation in forward genetic screening has been established in fish models early on (Driever et al., 1996; Wienholds et al., 2003). As a standard method, ENU (ethylnitrosourea) treatment introduced point mutations via base alkylation that give rise to single base mutations (often called ENU mutants) in zebrafish (Driever et al., 1996) and medaka (Loosli et al., 2000; Furutani-Seiki et al., 2004). Lacking efficient methods for targeted gene mutagenesis used to be a weak spot of zebrafish reverse genetics, but was recently overcome by the invention of zinc-finger nucleases (ZFN) (Doyon et al., 2008), transcription activator-like effector nucleases (TALENs) (Huang et al., 2011; Bedell et al., 2012), and the Clustered, Regularly Interspaced, Short Palindromic Repeat (CRISPR)/CRISPR-associated 9 (Cas9) technology (Hruscha et al., 2013; Hwang et al., 2013). CRISPR became the favorable genetic manipulation strategy given its ease to generate and assemble, and application in almost all eukaryotic cells. In addition to gene knockout/mutagenesis, CRISPR technology was further modified for generating knock-in/transgenic zebrafish carrying reporter/functional genes under endogenous gene regulation (Kimura et al., 2014). Like zebrafish, genome editing using TALEN (Ansai et al., 2013, 2014) and CRISPR/Cas9 (Ansai and Kinoshita, 2014) have been established in medaka. CRISPR/Cas9 mediated gene knock-in via NHEJ was used to generate transgenic and mutant medaka with a high germline transmission rate (Watakabe et al., 2018). In addition, the knock-in method using CRISPR was also applied to generate conditional knockout zebrafish by targeted insertion of loxP sites (Burg et al., 2018). Instead of NHEJ mediated knock-in which is error-prone, Wierson et al. (2020) have further developed knock-in method based on homology mediated end joining (HMEJ) repair for a more efficient and precise genome editing in zebrafish known as the GENEWELD method. Likewise, homology-directed repair (HDR) mediated knock-in strategies were also feasible in medaka (Murakami et al., 2017).

This concurrent development of technologies in zebrafish and medaka highlights the reciprocal nature of exchanging tools and methods between these model systems, facilitating the advancement of scientific research. Besides mutagenesis, genetic manipulation via stable transgenesis was first demonstrated in medaka (Ozato et al., 1986). Later, Kawakami and Shima (1999) identified the Tol2 transposon system in Medaka and adapted this system in zebrafish for transgenesis which revolutionized the field. To better recapitulate the endogenous gene expression patterns, insertion of BAC constructs by Tol2 transposase has been widely used in generating zebrafish reporter lines (Suster et al., 2011). Despite the fact that Tol2 was originally identified in medaka, it is more efficient in zebrafish (Kawakami, 2007). Therefore, another method for insertional transgenesis was developed in fish models using I-SceI meganuclease and greatly facilitated transgenesis in medaka (Thermes et al., 2002; Grabher and Wittbrodt, 2008).

As applications, spatial (e.g., specific tissue) and temporal regulation of ectopic gene expression can be achieved by combining transgenic lines generated by the above-mentioned methods (Tables 4, 5). For example, the Cre/lox system is widely used to perform reverse genetics, ectopic gene expression, and lineage tracing experiments in multicellular organisms, including zebrafish (Felker and Mosimann, 2016). To achieve spatial-temporal control, the Cre recombinase gene is fused with a human estrogen receptor (ER) domain and expressed under tissue-specific promoter, resulting in recombination of lox sequences in specific tissue upon estrogen stimulation (Metzger et al., 1995; Feil et al., 1996). Since then, there have been growing numbers of tissue-specific Cre lines generated in zebrafish (Jungke et al., 2013, 2015). Within the scope of tissue repair and regeneration, we have summarized a list of Cre driver and switch lines applied in previously described studies (Table 5). Additionally, robust co-expression of multiple genes following switch cassette can be accomplished by placing polycistronic ORFs separated by short viral 2A peptides (Provost et al., 2007). In contrast to zebrafish, there is way fewer medaka Cre/lox transgenic lines that we found and summarized with respect to the comprehensive list in zebrafish with available databases like CreZoo (Jungke et al., 2013, 2015). Fortunately, the advances of CRISPR technology allow genetic manipulation in both fish models and generate powerful tools for fate mapping and functional experiments (Liu et al., 2019). Overall, our review encompasses a portion of tools and strategies commonly used in zebrafish and medaka, which were extensively reviewed elsewhere for medaka (Kirchmaier et al., 2015) and zebrafish (Sassen and Köster, 2015), respectively.

Since zebrafish and medaka are routinely used for drug screening and validations, developing various routes of pharmaceutic administration is essential (Table 6). As these fish are tiny compared to mice, the development of administration methods requires further optimization to mimic the delivery route in mice.

Intraperitoneal (IP) injection was one of the first delivery methods introduced to deliver reagents in adult zebrafish and is adapted from veterinary practice in bigger fishes (Kinkel et al., 2010). The injection is performed using a 31G needle with a small injection volume (usually < 10–15 μl) into the abdominal cavity posterior to the pelvic girdle and midline to the pelvic fins in zebrafish (Kinkel et al., 2010). The reagents administrated by IP injection will distribute majorly to the spleen and liver within 72 h and later into the circulation, making it a favorable method for studying biochemical modulation in zebrafish over multiple injections or an extended period (Ruyra et al., 2014). IP is widely used for systemic administration of drugs, small-molecule inhibitors, nanoparticles, reagents in regeneration studies, including clodronate liposomes for ablating macrophages (de Preux Charles et al., 2016), poly I:C for immune-stimulation in medaka (Lai et al., 2017), mTOR-inhibitor rapamycin for autophagy inhibition (Chavez et al., 2020), IWR-1-endo for Wnt inhibition (Chen et al., 2009; Zhao et al., 2019), and tamoxifen for Cre-mediated recombination (Hans et al., 2009).

On the other hand, intravenous (IV) injection is effective for drug delivery in mice but has been technically challenging for zebrafish, due to the small vessel diameter and poor vasculature visibility. To improve vessel visibility, transparent casper fishes can be injected intravenously through their cardinal vein described in cancer research in zebrafish (Benjamin and Hynes, 2017). As alternative methods for direct access to the circulation, the intra-cardiac injection has been tested but showed high mortality (White et al., 2008), while the retro-orbital (RO) injection is suitable for delivering both reagents and cells into the blood circulation (Pugach et al., 2009; Simoes et al., 2020).

For more tissue/organ-localized delivery methods, intra-tissue injections may be applied. For example, intrathoracic (IT) injections were developed in zebrafish to test the effects of exogenous factors on adult heart regeneration, including nanoparticles encapsulated siRNAs (Xiao et al., 2018; Bise and Jazwinska, 2019). Cerebroventricular microinjection (CVMI) was also developed to deliver reagents to the adult zebrafish brain (Kizil and Brand, 2011). Intravitreal injections for the targeted delivery to the vitreous space of the retina (Fimbel et al., 2007) and intraspinal injection to the spinal cord were also developed in zebrafish (Wehner et al., 2018).

In contrast to invasive methods, which sometimes lead to tissue damage and mortality, a straightforward and convenient way for pharmaceutic delivery is immersion/incubation. Although immersion/incubation can be applied to both larvae and adults, it is more costly for incubating adults due to a large amount of reagent needed to reach the same dose. The biodistribution from immersion varies among reagents and different stages. In adults, the reagent is mainly intake through gills and ingestion, then absorbed/digested in the intestine and liver, which resembles oral uptake in the mice model (Ruyra et al., 2014). Examples of incubation experiments include tamoxifen treatment for Cre activation and Erbb2 inhibitor AG1478 in heart regeneration of both zebrafish larvae and adults (de Koning et al., 2015; Gemberling et al., 2015). Incubation of Alk5/4 inhibitor SB431542 has also been used in adult fin (Jaźwińska et al., 2007) and heart (Chablais and Jaźwińska, 2012b) to understand the dynamics of fibrosis in tissue regeneration.

Overall, based on the size and anatomic similarities between zebrafish and medaka, most of these delivery methods (Table 6) can be adapted in medaka and are regularly used in our laboratory as well as in other studies (Maekawa et al., 2016; Lai et al., 2017; Marín-Juez et al., 2019).

Due to the interest in exploring cellular and molecular mechanisms throughout the last few decades, both zebrafish (∼1,412 Mb) (Howe et al., 2013) and medaka genome (∼800 Mb) (Kasahara et al., 2007; Kobayashi and Takeda, 2008) have been sequenced and are publicly available in the databases, including Ensembl for zebrafish⁵ (see footnote 1) and medaka⁶ (Table 7). Like zebrafish, medaka has emerged as one of the most popular and influential animal models to investigate development and disease. At the genomic level, the regenerative capacity of zebrafish might rely on gene regulatory networks, which might be repressed in other non-regenerative animal models (Yang and Kang, 2019). Early findings in zebrafish suggest epigenetic modifications ranging from histone modifications to initiation of enhancer-induced activation of regenerative programs (Kang et al., 2016; Pfefferli and Jaźwińska, 2017). Growing evidence support the compatibility of zebrafish and medaka for comparative transcriptomic analyses (Tena et al., 2014; Lai et al., 2017). Moreover, with the advances of epigenetic profiling, medaka has also been explored for epigenetic changes associated with embryogenesis, development, and evolution (Nakamura et al., 2014; Tena et al., 2014; Ichikawa et al., 2017; Marletaz et al., 2018; Uesaka et al., 2019; Li et al., 2020). These properties may bring medaka as a new teleost model in epigenetics for comparative studies in tissue regeneration.

Among various teleost fishes that come in different shapes and sizes, zebrafish and medaka have a similar developmental process overall. However, medaka shows a slower pace and hatch at 9 days compared to 3 days in zebrafish. Considering the similarities and differences between zebrafish and medaka, inter-species blastula transplantation was explored to study the genetic developmental timing during organogenesis (Hong et al., 2012; Fuhrmann et al., 2020). Generating chimeric organisms named “Zebraka” or “Medrafish” involves cell transplantation at the blastula stage and ectopic formation of chimeric organs like retina from zebrafish donor cells in the medaka host. This approach helps researchers determine the transcriptional dynamics of retinal organogenesis and further state the existence of organ-intrinsic mechanisms independent of the development pace of the host (Fuhrmann et al., 2020). It will be intriguing to consider the differential regenerative capacity of each organ in these hybrid animals. For example, if the heart was contributed by zebrafish donor cells and grows in medaka, would it still be regenerative (and why)? Hybrid animals may help researchers dissect the intrinsic and extrinsic properties of tissue regeneration, and zebrafish and medaka chimeras may provide a unique opportunity for such study.

Similar ideas have been exploited in the intra-species blastula transplantation in embryonic and adoptive transfer in adult zebrafish to determine the cellular contribution of specific biological processes (cell-autonomous vs. non-cell-autonomous actions). The concept was recently adopted in a heart regeneration study where macrophages were isolated from zebrafish larvae donors and adoptively transferred into adult hosts. Larval macrophages were found to infiltrate the injured hearts and contribute to scar formation by directly secreting collagens in adults (Simoes et al., 2020). Similar strategies may pave the way for future research, in which zebrafish donor cells might be transferred in medaka host or vice-versa and determine the cellular contributions in tissue regeneration.