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eLife. 2017; 6: e22716.
PMCID: PMC5360449

Transient inflammatory response mediated by interleukin-1β is required for proper regeneration in zebrafish fin fold

Didier YR Stainier, Reviewing editor
Didier YR Stainier, Max Planck Institute for Heart and Lung Research, Germany;


Cellular responses to injury are crucial for complete tissue regeneration, but their underlying processes remain incompletely elucidated. We have previously reported that myeloid-defective zebrafish mutants display apoptosis of regenerative cells during fin fold regeneration. Here, we found that the apoptosis phenotype is induced by prolonged expression of interleukin 1 beta (il1b). Myeloid cells are considered to be the principal source of Il1b, but we show that epithelial cells express il1b in response to tissue injury and initiate the inflammatory response, and that its resolution by macrophages is necessary for survival of regenerative cells. We further show that Il1b plays an essential role in normal fin fold regeneration by regulating expression of regeneration-induced genes. Our study reveals that proper levels of Il1b signaling and tissue inflammation, which are tuned by macrophages, play a crucial role in tissue regeneration.


Research Organism: Zebrafish

eLife digest

Animals and other multicellular organisms all have at least some ability to regenerate lost or wounded tissues. Zebrafish are particularly good at this to the extent that they can replace damaged or lost body parts with exact replicas of the originals. In 2015, a team of researchers found that some mutant zebrafish that lack blood cells including immune cells are unable to regenerate lost tissues. This is because the cells that are primed to regenerate die instead, but it was not clear why this happens.

Many immune cells have roles in fighting infection and in responding to tissue damage.When a tissue is damaged, the area often becomes inflamed as white blood cells called macrophages flock to the damaged area to protect it from infection and remove damaged cells.

Hasegawa et al. – who include several researchers involved in the 2015 study – used genetic approaches to investigate the role of inflammation in tissue regeneration in zebrafish. The experiments show that several genes involved in inflammation – including one called interleukin 1b – were active over longer periods of time in the mutant fish compared with normal zebrafish. The gene produces a signal protein and this prolonged activity causes the primed regenerative cells to die. However, the cells can survive if interleukin 1b activity is quickly suppressed by macrophages. The experiments also show that, in order for tissues to regenerate properly, interleukin 1b needs to be active for only a short period of time.

The findings reveal that some inflammation is needed for tissues to regenerate, but that a more severe inflammatory response can block the process. A future challenge will be to identify the signals that macrophages produce to suppress inflammation to allow tissues to regenerate. These anti-inflammatory signals may have the potential to be used as drugs to cure chronic inflammatory diseases and boost tissue regeneration potential in humans.



Progress in regenerative medicine depends on unraveling the mechanisms underlying tissue and organ regeneration. The zebrafish is a powerful model species for investigation of regeneration mechanisms because it exhibits high regenerative capacity. Zebrafish can regenerate complex structures such as fins, heart, brain, retina, and other tissues (Gemberling et al., 2013). The caudal fin of zebrafish, in particular, has been used as a model for analyzing epimorphic regeneration, a type of regeneration in the urodele limb and fish fin (Poss et al., 2003). During epimorphic regeneration, two characteristic tissues, the wound epidermis and the blastema, are induced in response to tissue amputation, and their coordinated actions regulate cell proliferation and morphogenesis and thus lead to tissue regeneration. Studies conducted using the caudal fin regeneration model have identified numerous genes and molecular signaling pathways that are critical for fin regeneration (Yoshinari and Kawakami, 2011; Wehner and Weidinger, 2015).

In addition to the classical regeneration model developed using the adult caudal fin, a zebrafish larval fin fold model has been developed (Kawakami et al., 2004; Mateus et al., 2012), allowing identification of additional genes and signaling pathways required for regeneration (Mathew et al., 2009; Ishida et al., 2010; Yoo et al., 2012). Specifically, the fin fold regeneration model has facilitated genetic analysis of the regeneration mechanism by exploiting a number of mutant zebrafish resources (Rojas-Muñoz et al., 2009; Yoshinari et al., 2009), because several zebrafish lethal mutants survive beyond 7 days post fertilization (dpf)—an adequate period for assaying tissue regeneration—because of nutrient supply from the yolk.

Previously, we reported that zebrafish mutants, cloche (clo) (Stainier et al., 1995; Reischauer et al., 2016) and tal1/scl (Bussmann et al., 2007), showed a unique regenerative defect: these mutants could not regenerate their fin fold because of apoptosis of the regenerative cells (Hasegawa et al., 2015). Our analyses of the clo mutant revealed that a diffusible survival factor from myeloid-lineage cells is required to prevent apoptosis of the primed regenerative cells. That study suggested that the regenerative cells are sensitive to apoptosis during tissue regeneration, but the mechanism of this sensitization and the identity of the survival factor remain unknown.

Myeloid cells are widely recognized to play a role in protection against pathogens and microorganisms after tissue injury. Neutrophils are first recruited to sites of inflammation, and these cells encounter pathogens and phagocytose and kill microorganisms by producing reactive oxygen species and/or antibacterial proteins (Kolaczkowska and Kubes, 2013). Subsequently, macrophages infiltrate the inflammation site, produce pro- or anti-inflammatory cytokines, remove tissue debris, secrete growth factors, and support tissue restoration (Koh and DiPietro, 2011).

Several recent studies have used the zebrafish model and addressed the roles of myeloid cells in response to tissue injury: Li and coworkers (Li et al., 2012) showed that knockdown of macrophage differentiation delayed fin fold regeneration and resulted in formation of an abnormal fin fold featuring large vacuoles. Furthermore, Petrie and coworkers (Petrie et al., 2014) showed that genetic ablation of macrophages in adult zebrafish impaired fin outgrowth and often induced an abnormal fin. These studies suggest a crucial role for macrophages in tissue regeneration; however, the mechanism by which myeloid cells affect regeneration remains to be elucidated.

Here, we show that a pro-inflammatory cytokine gene, interleukin 1 beta (il1b), is aberrantly upregulated in the clo mutant, and that the resulting excessive and prolonged inflammation causes apoptosis of the regenerative cells. Moreover, although myeloid cells are widely regarded as the main cells that secrete pro-inflammatory cytokines such as Il1b and thereby promote inflammation, we demonstrate that epidermal cells surrounding the amputated tissue are the source of Il1b in both wild-type (WT) and clo larvae. Notably, il1b expression in the regenerating epidermis is usually quenched by the action of macrophages, and a lack of this anti-inflammatory effect of the macrophages results in apoptosis. Lastly, we show that Il1b signaling plays an indispensable role in normal tissue regeneration by activating expression of regeneration-induced genes. Thus, our study highlights the function of Il1b and the inflammatory response in tissue regeneration. A proper level of il1b expression and the inflammatory response, which are tuned by macrophages, are necessary for progression of tissue regeneration.


Prolonged il1b expression in the clo mutant during regeneration

To understand the molecular pathway leading to apoptosis of regenerative cells in the clo mutant, we performed a transcriptome analysis of amputated larval tail tissues of WT and clo mutant zebrafish at approximately 6 hr post amputation (hpa), just before the stage at which apoptosis is first detectable in the clo mutant (Hasegawa et al., 2015). Compared with WT larvae, the clo larvae showed marked upregulation of regeneration-induced genes such as junba, junbb, matrix metallopeptidase 9 (mmp9), fibronectin 1b (fn1b), and fgf20a (Yoshinari et al., 2009; Shibata et al., 2016) (Figure 1A and Supplementary file 1). In addition to these genes, expression of a group of genes including il1b and tumor necrosis factor-b (tnfb), which are involved in the inflammatory response, was also upregulated in the clo mutant during fin fold regeneration. Excluding enzymes and uncharacterized proteins, il1b was the most upregulated gene. il1b expression in the clo mutant was also confirmed using reverse transcription polymerase chain reaction (RT-PCR) analysis (Figure 1B). Notably, prostaglandin2a (ptgs2a), a downstream gene in Il1b signaling (Dinarello, 2009), was also upregulated in the clo mutant (Figure 1A), which clearly indicated that Il1b signaling was activated in the clo mutant.

Figure 1.
Prolonged activation of il1b expression in the clo mutant after fin fold amputation.

Next, we performed in situ hybridization (ISH) analysis of il1b to reveal its spatiotemporal expression during regeneration. At 3 hpa, strong il1b expression was observed in both WT and clo mutant larvae (Figure 1C and D). In WT larvae, il1b was expressed in a group of scattered cells that appeared to be myeloid cells (Figure 1C; arrowheads) and also expressed at the distal edge of the fin fold. After 3 hpa, il1b expression was rapidly diminished in WT larvae, and most larvae displayed very weak or no il1b expression at 6 hpa. In contrast to the expression in WT, il1b expression in the clo mutant was maintained at 6 hpa and was still detectable at 12 hpa, indicating a prolonged inflammatory reaction in the clo mutant.

Bacterial infection is recognized to trigger an inflammatory response that accompanies the induction of il1b expression (Dinarello, 2009). To investigate whether the same mechanism underlies the induction of il1b expression by fin fold amputation and bacterial infection, we injected zebrafish with Salmonella typhosa lipopolysaccharide (LPS), a reagent whose injection mimics bacterial infection (Lu et al., 2008). WT larvae injected with LPS displayed systemic induction of il1b in cells that appeared to be myeloid cells (Figure 1E). By contrast, the clo mutant larvae injected with LPS showed few il1b-positive cells, which indicated that tissue injury and bacterial infection induce il1b expression through distinct mechanisms.

il1b is expressed in epithelial cells during fin fold regeneration

The absence of myeloid cells in the clo mutant suggests that the il1b is expressed in non-myeloid cells. Accordingly, examination of tissue cryosections of the clo fin fold after il1b ISH analysis suggested that il1b was expressed in epidermal cells (Figure 2A).

Figure 2.
Spatiotemporal expression of il1b during fin fold regeneration.

To further investigate the identity of the il1b-expressing cells, we generated a transgenic (Tg) line, Tg(il1b:egfp), by introducing the egfp-nitroreductase fusion gene into the il1b locus of a bacterial artificial chromosome (BAC) clone, CH211-147H23. The larvae of Tg(il1b:egfp) showed faint EGFP expression in the entire epidermis and weak EGFP expression in the pectoral fins and at the end of the notochord (Figure 2—figure supplement 1A; arrowheads). After fin fold amputation, egfp expression was induced in a similar pattern to endogenous il1b expression (Figure 2B; Figure 2—figure supplement 1B), although EGFP fluorescence was detected at later stages of regeneration than il1b mRNA, probably as a result of the relatively longer half-life of the protein EGFP. LPS injection also induced EGFP fluorescence in myeloid cells (Figure 2—figure supplement 1C), which indicated that the il1b:egfp transgene recapitulated endogenous il1b expression in myeloid cells and also in the injured fin fold.

Next, using the Tg(il1b:egfp) reporter line, we confirmed that the EGFP-positive cells in the transgenic line mostly colocalized with the epithelial marker E-cadherin (Figure 2C), in both WT and the clo mutant; this finding indicated that il1b expression was principally induced in epithelial cells in response to tissue amputation. In addition to detecting il1b expression in the injured tissue, we detected il1b expression in WT larvae in migrating cells that are likely to be the myeloid cells (Video 1).

Video 1.

Time-lapse imaging of EGFP fluorescence in Tg(il1b:egfp) from 1 to 11 hpa.

Images were acquired once every 15 min. The epidermal cells at the injury site and the migrating cells, which are considered to be myeloid cells, expressed EGFP.


Il1b signaling and inflammation are responsible for apoptosis in the clo mutant

Excessive inflammation has been suggested to directly or indirectly induce apoptosis (Wallach et al., 2014). Therefore, because elevated expression of the pro-inflammatory cytokine Il1b leads to chronic inflammation (Dinarello, 2011), we suspected that excessive il1b expression in the clo mutant could be a cause of the apoptosis of regenerative cells.

First, we used the Tg(il1b:egfp) reporter line to examine the relationship between il1b-expressing cells and apoptotic cells. Although many of the il1b-expressing cells did not overlap with the apoptotic cells that were mainly distributed in the mesenchyme (Hasegawa et al., 2015), the cells lay in close apposition (Figure 3A).

Figure 3.
il1b knockdown or Dex treatment rescued the clo from apoptosis.

Next, to demonstrate the role of Il1b in the apoptosis occurring in the clo mutant, we knocked down il1b expression using an antisense morpholino oligonucleotide (MO). The il1b MO1 targeted to the splice sites (Nguyen-Chi et al., 2014) effectively inhibited il1b mRNA splicing (Figure 3B). Notably, il1b knockdown in the clo mutant resulted in a reduction in apoptosis of regenerative cells, whereas in the clo mutant injected with the standard control MO (std MO), no reduction in apoptosis was detected (Figure 3C and D). A similar anti-apoptosis effect of il1b knockdown was also obtained using another MO against il1b (Yan et al., 2014) (Figure 3—figure supplement 1).

To examine whether excessive inflammation is a cause of the apoptosis in the clo mutant, we used dexamethasone (Dex), a synthetic glucocorticoid that has been reported to suppress inflammation and il1b expression (Kern et al., 1988). Our results showed that Dex treatment abolished il1b expression in the clo mutant starting from an early stage of regeneration (Figure 3E), which indicated that the treatment effectively suppressed inflammation and il1b expression. In the Dex-treated clo larvae, the regenerative cells were rescued from apoptosis, much as in the il1b-knockdown larvae (Figure 3F and G). Collectively, these data support the notion that excessive Il1b signaling and inflammation in response to tissue injury represent a cause of apoptosis in the clo mutant.

Macrophages support survival of regenerative cells

We have previously shown that the presence of myeloid cells is essential for survival of regenerative cells (Hasegawa et al., 2015). Myeloid cells such as neutrophils and macrophages have been shown to transiently migrate to the injured fin fold (Li et al., 2012). Actually, we observed that the myeloid cell accumulation in the injured fin fold reached a maximum between 3 and 6 hpa (Figure 4A and B). The timing coincides with the stage at which the initial il1b expression immediately after wounding was downregulated (Figure 1C and D). This suggests that myeloid cell migration could play a role in attenuating il1b expression, although we cannot conclude that myeloid cell migration is necessary for il1b downregulation.

Figure 4.
Macrophages are responsible for survival of regenerative cells.

Myeloid cells comprise several different cell types, including neutrophils and macrophages, but the cell type required for the survival of regenerative cells remains to be determined. Here, we used csf3r, irf8, and spi1b (pu.1) MOs, which inhibited the differentiation of neutrophils, macrophages, and both types of cells, respectively, and examined whether the depletion of neutrophils or macrophages induces apoptosis of regenerative cells as in the clo mutant. The efficacy of csf3r knockdown was confirmed by Sudan Black staining of neutrophils and ISH analysis using the mpx probe. In both the csf3r morphants and the spi1b morphants, fewer neutrophils were detected than in the control morphants (Figure 4—figure supplement 1A,B,D). In the irf8 morphants, the expression of apoeb, a marker for microglia—the resident macrophages in the central nervous system—was drastically decreased (Figure 4—figure supplement 1C). The decrease of macrophages was also evident in the wounded fin fold as revealed by lcp1 ISH analysis (Figure 4—figure supplement 1E). None of these morphants showed any morphological abnormalities. These data indicated that the irf8 MO effectively inhibited macrophage differentiation.

To examine apoptosis, we performed TUNEL staining in the aforementioned morphants after fin fold amputation, and the results showed that irf8 morphants exhibited aberrant apoptosis of the regenerative cells, as did the spi1b morphants (Hasegawa et al., 2015) (Figure 4C). Importantly, similar numbers of TUNEL-positive cells were observed in the irf8 and the spi1b morphants (Figure 4D). In sharp contrast to the irf8 morphants, the csf3r morphants showed no increase in the apoptosis of regenerative cells. These results suggest that macrophages are responsible for supporting the survival of regenerative cells.

Macrophages attenuate il1b expression and inflammation

We next investigated the mechanism by which macrophages support the survival of regenerative cells. First, we examined il1b expression at 6 hpa in the spi1b, csf3r, and irf8 morphants. Whereas normal il1b expression was detected in the csf3r morphants, as in WT larvae, il1b expression remained elevated in the spi1b and irf8 morphants, as in the clo mutant (Figure 5A and B). This result suggests that macrophages attenuate il1b expression and thereby prevent apoptosis of regenerative cells in WT larvae. Furthermore, the apoptosis induced in the spi1b or irf8 morphants was rescued following il1b knockdown or Dex treatment (Figure 5C–E), which suggests that elevated il1b expression in the spi1b and irf8 morphants is responsible for the observed apoptosis.

Figure 5.
Prolonged il1b expression and apoptosis are induced by macrophage loss during fin fold regeneration.

il1b overexpression induces aberrant apoptosis and thereby suppresses fin fold regeneration

To further demonstrate that excessive il1b expression induces apoptosis, we generated and used the construct pTol2(hsp70l:mCherry-T2a-il1b), which we hereafter refer to as pTol2(hsp70l:il1b). The precise cleavage site of the zebrafish Il1b pro-peptide has not yet been identified (Vojtech et al., 2012), but based on comparing the zebrafish and human sequences, we estimated that the human IL-1β cleavage site (D116) corresponds to the threonine residue at the 124th amino acid (a.a.) position in zebrafish Il1b (Figure 6—figure supplement 1A). Furthermore, to obtain constitutive secretion of this Il1b protein lacking a canonical signal-peptide sequence, the Il1b sequence after a.a. 125 was fused with the human IL-1β receptor antagonist signal sequence (Wingren et al., 1996; Tu et al., 2008) and placed under the control of the promoter of heat shock protein 70l (Figure 6A). In the obtained Tg(hsp70l:il1b) line, mCherry fluorescence and il1b expression were observed after heat shock treatment (Figure 6B; Figure 6—figure supplement 1B), but not after stress caused by fin fold amputation (Figure 6—figure supplement 1C).

Figure 6.
il1b overexpression induces aberrant apoptosis.

When TUNEL staining was performed after heat shock induction at 2 days post amputation (dpa), several TUNEL-positive cells were detected in the injured fin fold (Figure 6C and D), which indicated that the primed regenerative cells were susceptible to the excess Il1b signal. Furthermore, systemic and aberrant apoptosis was also observed when il1b was overexpressed in WT embryo from 12 to 24 hr post fertilization (hpf), at which stage myeloid cells have not fully differentiated (Figure 6—figure supplement 1D). This indicated that the excessive Il1b signaling potentially induces apoptosis not only in fin fold but also in other parts of the body. In addition to inducing the aberrant apoptosis, il1b overexpression suppressed the extension of the fin fold (Figure 6E and F).

Normal inflammation mediated by Il1b is required for fin fold regeneration

Our data thus far suggest that excessive Il1b-mediated inflammation negatively affects the survival of regenerative cells and tissue regeneration in the clo mutant. However, inflammation has also been suggested to be essential for complete tissue regeneration (Mathew et al., 2007; King et al., 2012; Kyritsis et al., 2012). To examine the role of tissue inflammation and Il1b during normal fin fold regeneration, we used Dex and il1b MO1 and tested whether the inflammation mediated by Il1b affects normal fin fold regeneration.

To assess the role of inflammation in tissue regeneration, we administered Dex to WT larvae and assessed fin fold regeneration. The Dex-treated larvae displayed apparent retardation of regeneration at 3 dpa (Figure 7A and B) and a statistically significant reduction in cell proliferation in the distal fin fold region (Figure 7C and D). Notably, the expression of regeneration-induced genes such as junba (Yoshinari et al., 2009) and fgf20a (Whitehead et al., 2005; Shibata et al., 2016) was downregulated in the Dex-treated larvae (Figure 7E). These data suggest that inflammation is required for activation of regeneration-induced gene expression which is a prerequisite for normal regeneration.

Figure 7.
Il1b-mediated inflammation is required for normal regeneration.

Next, we used il1b knockdown in WT zebrafish to demonstrate the role of Il1b signaling in normal regeneration: il1b knockdown induced a similar phenotype to Dex treatment (Figure 7F–I), although the phenotype was slightly milder than that of the Dex-treated larvae, probably because of lower penetrance of the MO knockdown. Importantly, similar to the Dex-treated larvae, the il1b morphants showed attenuated expression of junba and fgf20a (Figure 7J). Collectively, these results suggest that the inflammatory reaction mediated by Il1b plays a necessary role in normal fin fold regeneration.

Lastly, we tested whether il1b overexpression induces a marked increase in expression of the regeneration-induced genes junba, junbb, and fn1b in uninjured tissue (Figure 7K). As expected, il1b overexpression induced the ectopic expression of fn1b, junba, and junbb, both in the uncut fin fold (Figure 7L) and in several other tissues such as the pectoral fins. Importantly, the apoptosis induced by il1b overexpression was only detectable at 2 days after multiple heat shocks (Figure 6C and D), possibly because of the anti-inflammatory function of macrophages in WT. But, induction of junba, junbb, and fn1b was observed at 12 hr after two heat shocks, a stage at which the apoptosis induced by il1b overexpression has not taken place (Figure 7—figure supplement 1), indicating that expression of the regeneration-induced genes was not caused by an indirect regenerative response to cell death. These data indicate that Il1b signaling is required tostimulate expression of regeneration-induced genes and regulate the initiation of fin fold regeneration.


It has been suggested that myeloid cells play crucial roles in the inflammatory responses of injured tissues. The pro-inflammatory cytokines (such as Il1b) produced by myeloid cells, including macrophages, induce early responses against infection or injury (Dinarello, 2009). Thus, myeloid cells are considered to trigger inflammation by providing pro-inflammatory cytokines. Here, we demonstrated that the pro-inflammatory cytokine Il1b is provided by epidermal cells in response to tissue injury. We showed that apoptosis occurred if the Il1b action at the injury site was prolonged, and that macrophages were responsible for attenuation of il1b expression and resolution of inflammation. Furthermore, our data suggest that normal il1b expression and inflammatory response are necessary in tissue regeneration to activate expression of regeneration-induced genes. Thus, our study has revealed that Il1b signaling and tissue inflammation act as a double-edged sword: they are required for regeneration, but in excess, they impair tissue regeneration (Figure 8).

Figure 8.
Schematic of the role of Il1b and macrophages during fin fold regeneration.

Myeloid cells are considered to be the principal producers of Il1b, although certain studies have reported il1b expression in melanoma cells (Okamoto et al., 2010) and human keratinocytes (Kupper et al., 1986). In this study, we showed that il1b is expressed by epidermal cells around the injury site. Notably, LPS injection induced il1b expression in myeloid cells, but not in the injured epidermal cells, suggesting that il1b induction by tissue injury occurs through a mechanism distinct from the mechanism underlying induction in myeloid cells following bacterial infection. Although it remains unknown how tissue injury stimulates il1b expression, a subject for future study, a substance released from disrupted cells (Rock and Kono, 2008) or a mechanical signal could be responsible for inducing il1b expression (Kanjanamekanant et al., 2013).

We also showed that in the clo mutant apoptosis is caused by prolonged il1b expression and inflammatory response. The precise mechanism by which Il1b induces apoptosis is unclear, but Tnf signaling could be a downstream mediator for triggering apoptosis. Tnfα signaling is known to stimulate apoptosis through activation of Caspase 8 (Sedger and McDermott, 2014). Tissue inflammation has been suggested to induce tnfα expression, and zebrafish tnfb, one of the zebrafish homologs of tnfα, was upregulated in the clo mutant (Figure 1A); therefore, Tnfα signaling potentially serves as a direct mediator for inducing apoptosis in the clo mutant. Alternatively, the clo apoptosis might occur because of ER stress signaling: Il1b administration into primary rat β-cells and MIN6 cells was reported to increase ER stress through intracellular release of Ca2+ and to induce apoptosis in a JNK-dependent manner (Eizirik and Mandrup-Poulsen, 2001; Wang et al., 2009). Thus, excessive ER stress evoked by Il1b around the injury site could induce apoptosis in the clo mutant.

In this study, we further demonstrated that macrophages play a crucial role in fin fold regeneration by attenuating il1b expression and supporting survival of regenerative cells. Recent studies have shown that knockdown of macrophage differentiation delayed fin fold regeneration and resulted in an abnormal fin fold featuring large vacuoles (Li et al., 2012), and genetic ablation of macrophages in adult zebrafish was demonstrated to affect fin ray patterning and fin growth (Petrie et al., 2014). These studies suggest that macrophages play a critical role in tissue regeneration, but the precise function of the cells remains unknown. Our study has, for the first time, elucidated a role of macrophages in attenuating il1b expression and inflammation during tissue regeneration. In accordance with our observation, macrophage-depleted mice and salamanders were found to display elevated il1b expression (Goren et al., 2009; Godwin et al., 2013), which suggests that the macrophage function of attenuating il1b transcription is conserved in other vertebrate species.

Our observation of macrophage accumulation in the injured fin fold suggests that the macrophage migration could play a role in attenuating il1b expression and quenching inflammation. However, we cannot conclude that the macrophage migration is necessary for attenuating the il1b expression, because our preceding study showed that an activity that rescues the clo apoptosis exists in the WT body extract prior to tissue amputation. We previously suggested that the effect of myeloid cells is mediated by a factor that is diffusible and heat-stable (Hasegawa et al., 2015), but its identity remains unknown. The factor could be an anti-inflammatory protein/molecule such as IL-10, Tgf-β, or a lipid mediator (Serhan et al., 2008; Ortega-Gómez et al., 2013). It is speculated that signals such as nFkB and/or STAT3, which are activated by tissue injury, are involved in inducing il1b expression (Fang et al., 2013; Ogryzko et al., 2014; Karra et al., 2015). The anti-inflammatory molecules bind to cell surface receptors to activate intracellular pathways and antagonize the nFkB and/or STAT3 signals to attenuate il1b transcription (Yu et al., 2009; Saraiva and O'Garra, 2010; Liao et al., 2012).

Most importantly, our study suggests that inflammation mediated by Il1b is also necessary for normal fin regeneration. Previously, we showed that genes such as fn1b, junba, and junbb were induced in response to tissue injury (Yoshinari et al., 2009), but the upstream mechanism that regulates the expression of these genes remains unknown. Here, we have demonstrated that Il1b signaling is necessary and sufficient for triggering expression of regeneration-induced genes. Although the primary factors involved in tissue regeneration, tentatively named alarmins (Bianchi, 2007), remain to be identified, Il1b could represent one of the key upstream mediators for initiating expression of regeneration-induced genes and advancing tissue regeneration.

Given that macrophages are required for resolving inflammation, an impairment or reduction of this function will lead to the diseases that accompany chronic inflammation, such as rheumatoid arthritis, inflammatory bowel disease, and auto-inflammatory diseases. Therefore, elucidation of the regulatory mechanism of the Il1b signal and the action of anti-inflammatory substances released from macrophages will enhance our understanding of the pathogenic processes of chronic-inflammation diseases and lead to development of suitable therapeutics.

Materials and methods

Fish husbandry and fin amputation

Zebrafish of a WT line, which has been maintained in our facility for >10 years through inbreeding, were housed in a recirculating system in a 14 hr day/10 hr night cycle at 28.5°C. When necessary, larvae were incubated in egg water (0.06% artificial marine salt, 0.0002% methylene blue) containing 0.003% phenylthiourea to prevent pigment formation. The zebrafish mutant strain clom39 (RRID:ZFIN_ZDB-ALT-980203-381), the fgf20a enhancer-trap line HGn21A (Shibata et al., 2016), the BAC Tg(il1b:egfp) line, and Tg(hsp70l:il1b) were used. The clom39 mutant was genotyped as previously described (Hasegawa et al., 2015). Zebrafish were subject to heat-shock induction at 38°C for 1 hr in a small water bath and then cooled to 28.5°C over time. Fin fold fin amputation was performed as previously described (Kawakami et al., 2004). Briefly, zebrafish larvae at 2 dpf were anesthetized with 0.04% 3-amino benzoic acid ethylester (tricaine) in egg water and their fin fold was amputated using a surgical razor blade. For reproducible quantification, the fin fold was carefully amputated at sites just posterior to the notochords. For quantification of fin fold regeneration in Figure 6F, Figure 7B and G, the length from the notochord end to the posterior tip of fin fold was measured.

RNA sequencing analysis

Larval posterior tissues (~1 mm from the distal end, approximately the tissue posterior to the yolk extension) from uncut WT, amputated WT, and amputated clo mutant, which were obtained from more than five independent batches of incrosses of clo heterozygotes, were collected on dry ice and stored at −80°C. Total RNAs from approximately 500 posterior tissue samples were extracted using TRIzol (Thermo Fisher Scientific, Waltham, MA) according to the manufacturer’s instructions. RNA sequencing analysis was performed by Dragon Genomics (Kyoto, Japan). Briefly, purified and fragmented PolyA+ RNAs were prepared from 3 μg each of the total RNAs using the TruSeq RNA sample preparation kit (Illumina, San Diego, CA), and the respective cDNAs were synthesized according to the manufacturer’s instructions. The cDNAs fused with an adaptor on both ends were PCR-amplified (15 cycles) and used for sequencing analysis (Illumina HiSeq2000). The obtained sequence reads (60927373 reads for WT uncut, 59284653 reads for amputated WT at 6 hpa, 58507894 reads for amputated clo mutant at 6 hpa) were aligned to the zebrafish reference sequence using Bowtie software (version 0.12.7; RRID:SCR_005476) (Langmead et al., 2009). The normalized expression level of the respective genes, RPKM (reads per kilobase of exon model per million mapped reads), was calculated using ERANGE software (version 3.2; RRID:SCR_005240) (Mortazavi et al., 2008).

MO injection

MOs (Gene Tools, Philomath, OR) were dissolved in Danieau solution. Fertilized zebrafish eggs were dechorionated by incubating them with 2% pronase (Roche) and then microinjected at the 1–2 cell stages. The following MOs were used in this study:

irf8 MO: 5ʹ-AATGTTTCGCTTACTTTGAAAATGG-3ʹ (Hall et al., 2014)

csf3r MO1: 5ʹ-ATTCAAGCACATACTCAC-TTCCATT-3ʹ (Hall et al., 2014)

csf3r MO2: 5ʹ- GAACTGGCGGATCTGTAAAGACAAA −3ʹ (Halloum et al., 2016)

spi1b MO: 5ʹ-GATATACTGATACTCCATTGGTGGT-3ʹ (Rhodes et al., 2005)

il1b MO1: 5ʹ-CCCACAAACTGCA-AAATATCAGCTT-3ʹ (Nguyen-Chi et al., 2014)

il1b MO2: 5ʹ- AAACGTAAAATAACTCACCATTGCA −3ʹ (Yan et al., 2014)


The csf3r MO2 (0.25 mM) and the rest of MOs (1 mM) were injected at approximately 0.5 nl per egg. Under these conditions, no apparent side effect was observed. Injected embryos were incubated in 0.3× Niu-Twitty solution.

RT-PCR analysis

Total RNAs were extracted from larval tail tissues (200 for each) posterior to the yolk extension using TRIzol and further purified using the RNeasy kit (Qiagen, Venlo, Netherlands). Among the purified total RNAs, aliquots that corresponded to 90 larval tails were used for cDNA synthesis. The cDNAs were synthesized using the Thermoscript RT-PCR kit (Thermo Fisher Scientific) with random hexamers as the primer. The synthesized cDNAs were diluted to 0.1 μg/μl and stored at −30°C. PCR was performed according to a standard procedure using Paq5000 DNA polymerase (Agilent Technologies, Santa Clara, CA), and the products were analyzed using 2% agarose gels. The following primers were used for PCR:





Whole-mount ISH analysis

Whole-mount ISH analysis was performed as described previously (Hasegawa et al., 2015), except that 5% polyvinyl alcohol was included in the buffer during the color reaction. After detection of the ISH signal, the samples were fixed with 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) for color preservation, equilibrated with 80% glycerol, and then mounted on glass slides and photographed. The antisense junba, junbb, fn1b, apoeb, mpx, and lcp1 (l-plastin) probes used here were described previously (Yoshinari et al., 2009; Hall et al., 2012). The egfp probe was synthesized from the construct pCS2-egfp, which contained the full-length egfp coding sequence. The il1b probe was directly synthesized from the il1b PCR product using the T7 promoter (underlined in the primer sequence) (Thisse and Thisse, 2008).



For preparing tissue sections, samples were incubated in 20% sucrose in PBS for overnight at 4°C, embedded in Tissue-Tek compound (Miles) and 16 μm sections were obtained using a cryostat.

LPS injection

S. typhosa LPS (Sigma–Aldrich, St. Louis, MO) was dissolved in water at 10 mg/ml and stored in aliquots at −30°C. A solution containing LPS (5 mg/ml) and phenol red (0.5%) was injected at 0.5 nl per larva into the pericardial cavity of anesthetized larvae at 2 dpf.

Generation of Tg lines

The construct for generating Tg(hsp70l:mCherry-2a-il1b) was prepared by replacing the creERt2 sequence of pT2(hsp70l:mCherry-2a-creERt2) (Yoshinari et al., 2012) with the il1b cassette. The il1b cassette was prepared by PCR-amplifying the il1b sequence a.a. 125–273. The signal sequence of the human IL-1β receptor antagonist (Wingren et al., 1996) and the HA tag were fused at the N-terminus and C-terminus, respectively (underlined in the primer sequences).



The construct was injected into fertilized eggs. To identify germ-line transmission, the F0 fish were crossed with each other or with WT, and the embryos produced were screened for mCherry fluorescence after heat shock.

To generate Tg(il1b:egfp), the iTol2 cassette was first introduced into the BAC clone CH211-147H23 using BAC recombineering (Suster et al., 2011). Next, the egfp-nitroreductase cassette (Grohmann et al., 2009) was introduced into the site of the initiation codon of il1b. Lastly, a mixture of purified BAC DNA (125 ng/μl) and the transposase mRNA (25 ng/μl) was injected into 1-cell-stage zebrafish embryos (1 nl/embryo). The Tg was screened for EGFP expression in the lens, which is driven by the crystalline alpha promoter, and further tested for EGFP induction in response to fin fold amputation.

Whole-mount immunohistochemistry

Zebrafish larvae were fixed with 4% PFA in PBS for 2 hr at room temperature (RT) or overnight at 4°C, washed thrice with PBS containing 0.1% Triton X-100 (PBTx), dehydrated with methanol, and stored at −30°C. Samples were rehydrated with PBTx and then incubated (overnight, 4°C) with anti-E-cadherin (1:1000; BD Biosciences, Franklin Lakes, NJ; RRID:AB_397580) and anti-EGFP (1:1000; Nacalai Tesque, Kyoto, Japan; RRID:AB_10013361) antibodies in blocking buffer (5% serum and 0.2% bovine serum albumin in PBTx). After extensive washing with PBTx at RT, the samples were incubated (overnight, 4°C) with anti-mouse Alexa Fluor 568 and anti-rat Alexa Fluor 488 antibodies (both 1:1000; Invitrogen). After washing with PBTx, the tail regions were isolated and mounted in 80% glycerol containing 2.5% 1,4-diazabicyclo[2.2.2]octane (Nacalai Tesque) as an anti-fading reagent. Fluorescence images were acquired using a confocal microscope (FV-1000, Olympus, Tokyo, Japan).

Time-lapse analysis of Tg(il1b:egfp)

Zebrafish larvae of the BAC Tg(il1b:egfp) line at 1 hpa were placed on a 2% agarose-gel stage in egg water and embedded in 0.7% low-melting agarose. Time-lapse images were acquired once every 15 min using the confocal microscope equipped with a 20× water-immersion objective lens; the acquired images were z-stacks containing 20 optical slices.

TUNEL staining

Larvae were fixed with 4% PFA for 2 hr at RT or overnight at 4°C, dehydrated with methanol, and stored at −30°C. Apoptosis was examined using an in situ apoptosis detection kit (Roche, Basel, Switzerland). Briefly, samples were rehydrated with PBTx, treated with 10 μg/ml Proteinase K in PBTx (5 min, RT), washed with PBTx, and refixed with 4% PFA in PBS (20 min). The samples were further incubated (15 min, on ice) in a freshly prepared 0.1% sodium citrate buffer containing 0.1% Triton X-100, washed with PBTx, and reacted with the TUNEL reaction mixture at 37°C for 1.5 hr. The reaction was terminated by washing with PBTx. The samples were mounted in 80% glycerol and fluorescence images were acquired using confocal microscopy. The TUNEL-positive cells were quantified by counting the number of cells in the area posterior to the notochords.

Chemical treatment

Dex (D-2915; Sigma) was dissolved in dimethyl sulfoxide (DMSO) at 100 mM and stored at −30°C. The Dex solution was diluted to 100 μM with egg water and administered to zebrafish larvae at least 1 hr before fin fold amputation.

Sudan Black B staining

Larvae were fixed with 4% PFA for 1 hr at RT, rinsed with PBS, and incubated in 0.03% Sudan black B (Sigma). After extensive washing with 70% ethanol and rehydration with PBS containing 0.1% Tween 20 (PBT), samples were mounted in 80% glycerol.

BrdU staining

Proliferating cells were labeled with 5-bromo-2-deoxyuridine (BrdU, 5 mM) during 0–12 hpa. The labeled larvae were fixed with 4% PFA for 2 hr at RT or overnight at 4°C, dehydrated with methanol, and stored at −30°C. Immunochemical detection was performed as described (Yoshinari et al., 2009) and BrdU-positive cells were quantified from the acquired confocal images. Similar to the quantification of TUNEL-positive cells, the BrdU-positive cells were quantified by counting the number of cells in the area posterior to the notochords.


Most assessments of phenotypes and expression patterns were replicated in at least two independent experiments with comparable results. Larvae were collected from independent crosses, and experimental processing (injection, heat shock, and/or staining) was carried out on independent occasions. Exceptions to this include data presented in Figure 1E, Figure 7L, Figure 2—figure supplement 1, and Figure 6—figure supplement 1B–D. In each of these cases, multiple larvae were processed, and the obtained phenotypes were the same in all or most cases. The n is reported within the respective figures.

Statistical analysis

Data are presented as means ± SEM. Statistical analyses were performed using Microsoft Excel 2013. For normally distributed data, differences were analyzed using Student’s t tests; p<0.05 was considered to be statistically significant.


We thank members of the Kawakami lab. This work was supported by grants from the Koyanagi Foundation and a Grant-in-Aid for Scientific Research (C) to A Kawakami, by the Marsden Fund grant from the Royal Society of New Zealand to C. J Hall, and by the NIG-JOINT grant from the National Institute of Genetics to A Kawakami and K Kawakami, and by the National BioResource Project from Japan Agency for Medical Research and Development (AMED) to K Kawakami. TH was supported by a fellowship from the Education Academy of Computational Life Science (ACLS) at Tokyo Institute of Technology.

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Funding Information

This paper was supported by the following grants:

  • Society for the Promotion of Science Grant-in-Aid for Scientific Research (C) to Atsushi Kawakami.
  • Agency for Medical Research and Development National BioResource Project to Koichi Kawakami.
  • Society of New Zealand Marsden Fund to Christopher J Hall.

Additional information

Competing interests

The authors declare that no competing interests exist.

Author contributions

TH, Conceptualization, Formal analysis, Investigation, Writing—original draft, Writing—review and editing.

CJH, Resources, Data curation, Supervision, Funding acquisition, Investigation, Methodology.

PSC, Data curation, Supervision, Funding acquisition.

GA, Resources, Investigation, Methodology.

KK, Resources, Funding acquisition, Methodology.

AKu, Supervision, Project administration.

AKa, Conceptualization, Resources, Data curation, Formal analysis, Supervision, Funding acquisition, Investigation, Methodology, Writing—original draft, Project administration, Writing—review and editing.


Animal experimentation: This study was performed in strict accordance with the recommendations in the Act on Welfare and Management of Animals in Japan and the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All of the animals were handled according to the Animal Research Guidelines at Tokyo Institute of Technology. The protocol was approved by the Committee on the Ethics of Animal Experiments of the Tokyo Institute of Technology. All surgery was performed under tricaine (3-aminobenzoic acid ethyl ester) anesthesia, and every effort was made to minimize suffering.

Additional files


Supplementary file 1.

The list of transcripts that are upregulated or downregulated in the amputated fin fold of the clo mutant.

The table of upregulated genes shows the list of transcripts whose final RPKMs of the amputated clo mutant at 6 hpa are more than two times than those of the amputated WT. The table of downregulated genes shows the list of top 100 transcripts whose final RPKMs of the amputated clo mutant at 6 hpa are downregulated compared with those of amputated WT. Transcripts with low expression in the clo mutant (final RPKM <4) were excluded. Final RPKM, the reads per kilobase of exon model per million mapped reads. Transcripts that are shown in Figure 1A are highlighted.



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2017; 6: e22716.

Decision letter

Didier YR Stainier, Reviewing editor
Didier YR Stainier, Max Planck Institute for Heart and Lung Research, Germany;

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Transient inflammatory response mediated by interleukin-1β is required for proper regeneration in zebrafish fin fold" for consideration by eLife. Your article has been favorably evaluated by Robb Krumlauf (Senior Editor) and three reviewers, one of whom is a member of our Board of Reviewing Editors. The reviewers have opted to remain anonymous.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission


This interesting paper from the Kawakami lab follows up on their earlier publication reporting that fin regeneration fails to occur in zebrafish cloche mutants due to the lack of myeloid cells.

In this new study, they show that il1b is a likely factor involved in the lack of fin regeneration in cloche mutants using both loss- and gain-of-function analyses.

Interestingly, il1b appears to be expressed by epithelial cells, not myeloid cells, and macrophages appear to be necessary for the resolution of inflammation/end of il1b expression. Prolonged il1b expression leads to apoptosis.

Essential revisions:

1) When do macrophages arrive near the wound, and how does that relate with the down-regulation of il1b expression? Defining macrophage as a key cell type mediating cell survival during fin fold regeneration is one of the major conclusions of this paper. Thus, the characterization of macrophages in this mechanism would need to be elaborated more. Is there any evidence that macrophage infiltration in the vicinity of il1b-positive epithelial cells occurs after the initial priming stage (Figure 8C)? Is it possible that they infiltrate immediately after the injury and change their function afterwards to suppress il1b signals? Or, is it possible that macrophages are not residing in the regenerating area and mediate the suppressive function remotely?

These questions could be addressed by using mpeg1 reporter fish, its reporter construct injection, or in situ hybridization.

2) Overexpression of il1b induced pro-regenerative gene expression in uninjured fins (Figure 7L). This result may also be interpreted as an indirect regenerative response to cell death induced in uninured fins with excessive il1b (Figure 6—figure supplement 1C), not as evidence for the direct role of il1b in regeneration priming.

One way to address this question is by doing TUNEL staining at a time point when the regenerative genes are induced in the uninjured, il1b-overexpressing fin to determine whether these genes are expressed unrelated to or earlier than Il1b-mediated cell death induction.

Essential revisions:

1) When do macrophages arrive near the wound, and how does that relate with the down-regulation of il1b expression? Defining macrophage as a key cell type mediating cell survival during fin fold regeneration is one of the major conclusions of this paper. Thus, the characterization of macrophages in this mechanism would need to be elaborated more. Is there any evidence that macrophage infiltration in the vicinity of il1b-positive epithelial cells occurs after the initial priming stage (Figure 8C)? Is it possible that they infiltrate immediately after the injury and change their function afterwards to suppress il1b signals? Or, is it possible that macrophages are not residing in the regenerating area and mediate the suppressive function remotely?

These questions could be addressed by using mpeg1 reporter fish, its reporter construct injection, or in situ hybridization.

Migration of myeloid cells in response to tissue injury has been described in preceding studies using transgenic zebrafish and ISH analysis (Li et al., 2012; Yohisnari et al., 2009). According to the reviewer’s suggestion, we further examined temporal changes of neutrophil and macrophage accumulation, respectively, in the injured fin fold. We observed that both of neutrophil and macrophage increased until 3 hpa, reached at a maximum number between 3-6 hpa in the vicinity of il1b-expressing site, and then decreased thereafter. The timing coincides with the stage when the initial il1b expression in WT was downregulated, suggesting a possibility that myeloid cell migration could play a role for attenuating il1b expression. However, we cannot conclude that it is necessary for attenuating il1b expression. Because, we have previously shown that an anti-apoptosis activity exists in the WT body fluid irrespective of tissue injury (Hasegawa et al., 2015).

Figure 4A and B: We added the ISH analyses of neutrophil and macrophage accumulation in amputated fin fold.

Subsection “Macrophages support the survival of regenerative cells”, first paragraph: We added a description of myeloid cell accumulation in amputated fin fold in the Results section.

Discussion, fifth paragraph: We also added a discussion regarding the significance of myeloid cell migration to injured site.

Figure 8: The illustration in Figure 8C was revised, because we cannot conclude that macrophage infiltration in the vicinity of il1b-positive epithelial cells is necessary for attenuating il1b expression.

2) Overexpression of il1b induced pro-regenerative gene expression in uninjured fins (Figure 7L). This result may also be interpreted as an indirect regenerative response to cell death induced in uninured fins with excessive il1b (Figure 6—figure supplement 1C), not as evidence for the direct role of il1b in regeneration priming.

One way to address this question is by doing TUNEL staining at a time point when the regenerative genes are induced in the uninjured, il1b-overexpressing fin to determine whether these genes are expressed unrelated to or earlier than il1b-mediated cell death induction.

In the used Tg(hsp70l:il1b), multiple heat shock induction (5 times taking 2 days) was required before detecting the aberrant apoptosis in uncut WT fin fold. This is probably because the macrophages in WT suppress the inflammatory effect of Il1b overexpression. The induction of regenerative genes shown in Figure 7L was observed at 6 hpa, a stage before the apoptosis by il1b overexpression was evident. Indeed, we confirmed that apoptosis was not detected in the uncut fin fold after two HS inductions within 2 days.

Figure 7—figure supplement 1: We added new data.

Subsection “Normal inflammation mediated by Il1b is required for fin fold regeneration”, last paragraph: We added sentences describing that the expression of the regeneration-induced genes was not due to an indirect regenerative response to cell death.

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