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The regenerative capacities of vertebrate tissues/organs tend to decrease after repeated injury or when the animals become older. This is likely due to reduced progenitor cell proliferation and differentiation (Janzen et al.,2006; Collins et al.,2007; Nave,2008; Kirschner et al.,2010). While amphibians have long been the central characters employed in studies on tissue or organ regeneration (Brockes,1997; Beck et al.,2009; Contreras et al.,2009; Kragl et al.,2009; Calve et al.,2010), the zebrafish (Danio rerio) have recently emerged as a new vertebrate model for genetic studies of tissue/organ regeneration. Like amphibians, zebrafish exhibit an enhanced capability of regenerating adult tissues, which include retina, spinal cord, kidney, heart, and fin (Poss et al.,2000a,b,2002a,b; Nechiporuk and Keating,2002; Nechiporuk et al.,2003; Jazwinska et al.,2007; Schoenebeck et al.,2007; Tsai et al.,2007; Qin et al.,2009; Jopling et al.,2010; Thummel et al.,2010). Fin regeneration is a particularly efficient model for studying tissue regeneration. Although the fins may vary in shape, length, and color, they are built by identical skeletal elements (Becerra et al.,1983; Geraudie et al.,1995; Johnson and Bennett,1999, Kawakami et al.,2004; Lee et al.,2005; Murciano et al.,2007; Huang et al.,2009; Kizil et al.,2009). In response to injury, major fin structures may regenerate very rapidly in zebrafish. For example, within a few hours post fin amputation, epidermal cells migrate and accumulate, mesenchymal cells reorganize and proliferate, and new segments progressively add to the distal end of the regenerating fin until the original length of the fin is achieved (Geraudie and Singer,1992; Johnson and Weston,1995; Kawakami et al.,2004; Jazwinska et al.,2007; Borday et al.,2001; Murciano et al.,2007; Tsai et al.,2007).
Genes and cell-signaling pathways that are involved in zebrafish fin regeneration have been previously characterized (Laforest et al.,1998; Tawk et al.,2000; Quint et al.,2002; Padhi et al.,2004; Smith et al.,2006; Jazwinska et al.,2007; Stoick-Cooper et al.,2007a,b; Wills et al.,2008; Yin et al.,2008; Chablais and Jazwinska,2010). For example, Whitehead et al. (2005) reported that the expression of fgf20a, a gene that encodes a member of the fibroblast growth factor family, plays important roles in the initiation of fin regeneration, and that decreases in fgf20a expression result in inhibition of fin regeneration. Akimenko et al. (1995) and Nechiporuk and Keating (2002) demonstrated that the expression of msxb, a gene that encodes a transcriptional repressor that is commonly found in proliferating cells, is also required for fin blastema formation after fin injury. During regeneration, msxb is expressed in both proliferating and non-proliferating blastema cells. Inhibition of msxb expression results in decreased fin outgrowth (Thummel et al.,2006).
While the cellular events and molecular mechanisms of zebrafish fin regeneration have been well described, the potential of fin regeneration remains to be further studied (i.e., regeneration capability in response to repeated injuries, and regeneration in aged animals). In this research, we examined regeneration of the caudal fin in adult zebrafish that received multiple injuries at different ages. We measured caudal fin outgrowth at different times post-amputation and quantified the expression of regeneration marker genes (e.g., msxb, fgf20a, and bmp2b). The results provide evidence that the zebrafish has unlimited potential for regenerating the injured caudal fin.
The Expression of msxb and fgf20a in Regenerating Caudal Fins
The expression of regeneration marker genes, such as msxb and fgf20a, in regenerating fins has been previously characterized (Whitehead et al.,2005; Wills et al.,2008; Nechiporuk et al.,2003). In this study, we examined the levels of msxb and fgf20a expression in zebrafish caudal fin at different times post-amputation. Both msxb and fgf20a were expressed in the regenerating caudal fin (Fig. 1A). Upon injury, the expression of msxb and fgf20a was up-regulated. The expression of msxb increased rather slowly, reaching the highest level approximately 48 hr post-amputation (hpa), at which time the expression of msxb was approximately 6-fold higher than its expression measured in uninjured control fin tissue (designated 0 hpa) (Fig. 1B). After 48 hpa, the expression of msxb gradually decreased. Within 24 hours, the expression of msxb decreased to levels that were approximately three quarters of the peak expression levels. The expression of msxb remained low during late regeneration stages (up to 96 hpa examined; Fig. 1B).
The expression of fgf20a rapidly increased in response to fin amputation. By 24 hpa, the expression of fgf20a reached to levels that were approximately 15-fold higher than the expression measured in the uninjured caudal fin (0 hpa). Thereafter, the expression began to decrease. By 48 hpa, the expression of fgf20a decreased to levels that were approximately one quarter of the peak expression levels. The expression of fgf20a remained low during late regeneration (up to 96 hpa, Fig. 1C).
The Potential for Fin Regeneration Did Not Decline With Age
To determine whether fin regeneration correlates with age, we examined caudal fin regeneration in zebrafish at different ages (between 4 and 28 months old). The fish received fin amputation, and were allowed to recover at 33°C. We examined fin regeneration (tissue outgrowth and the expression of regeneration marker genes) at different times post-amputation.
We did not observe statistical differences in the initial tissue outgrowth or in the expression of msxb and fgf20a between young and old animals. At 48 hpa, caudal fin outgrowth was 0.90 ± 0.08 mm in animals at 4 months of age. Similar fin outgrowth was seen in zebrafish at 12, 18, and 28 months (Fig. 2A). At 48 hpa, the expression of msxb and fgf20a was up-regulated by 4–6-fold, respectively, in 4-month-old animals. Similar up-regulation of msxb and fgf20a expression was seen in animals at 12, 18, and 28 months (Fig. 2B, C).
We examined caudal fin outgrowth in late regeneration stages (4, 7, 20, and 30 days post-amputation; dpa). Similar fin outgrowth was observed in young and old animals when examined at 4 and 7 dpa. At 4dpa, for example, the injured caudal fin grew to about 40% of its original length (measured before amputation) in both young and old animals. By 7dpa, the regenerating fin grew to about 60% of the original length. However, slight differences in fin outgrowth were detected when measured at 20 and 30 dpa. At 20 dpa, the fin grew to about 90–93% of its original length in 4- and 12-month-old animals, and only 85% of its original length in 18-month-old animals (Fig. 3). At 30 dpa, the caudal fin regenerated to 95–98% of its original length in 4- and 12-month-old animals, but remained less than 90% in 18-month-old animals (Fig. 3). The data suggest that the ability to fully regenerate the injured caudal fin is slightly decreased in aged animals.
The Potential for Fin Regeneration Did Not Decline After Repeated Injury
We examined caudal fin regeneration in zebrafish that received repeated amputations. In these experiments, we amputated the caudal fin multiple times at 48-hr intervals. Re-amputation surgeries were performed at the original site where the first amputation took place. After each amputation, the fish were allowed to recover at 33°C. We examined fin regeneration, e.g., tissue growth and the expression of regeneration marker genes, at different times post-amputation. To ensure that we completely removed the blastema (formed after the previous amputation), we monitored the expression of msxb by in situ hybridization and qRT-PCR (Fig. 4A, B). In the control tissue (immediately after amputation) and in the re-amputated tissue (in which the blastema was removed), we observed no msxb expression. In tissues in which the blastema remained (formed after the previous amputation), msxb expression was detected.
In zebrafish that received only one amputation, at 48 hpa the caudal fin regenerated by 0.90 ± 0.08 mm. Similar fin outgrowth was seen in animals that received multiple amputations (2–9 times). We did not detect statistical differences in fin outgrowth after single or multiple amputations (Fig. 5A). Up-regulation of regeneration marker genes (msxb and fgf20a) was seen in all animals examined, regardless of whether they received single or multiple amputations. In all cases, at 48 hpa the expression of msxb and fgf20a increased by 4–6-fold, respectively, in comparison to expression measured in control tissues (0 hpa) (Fig. 5B, C).
We examined caudal fin regeneration in late regeneration stages (e.g., at 7 dpa) after single or multiple amputations (up to 6 times). The experiments were performed in zebrafish at different ages (4 and 12 months old). At 7 dpa, the regenerating caudal fin grew to approximately 60% of its original length in all fish examined, regardless of whether they received single or multiple amputations. No differences were seen in fin outgrowth in 4- and 12-month-old animals (Fig. 6). We quantified the expression of another regeneration marker gene, bmp2b, which is up-regulated during fin outgrowth (Quint et al.,2002; Smith et al.,2006). At 7 dpa, we observed no statistical differences in bmp2b expression in response to single or repeated amputations or in fish at different ages (Fig. 6).
Tissue regeneration has been previously studied using both amphibian and fish models (Johnson and Weston,1995; Geraudie and Singer,1992; Brockes,1997; Jazwinska et al.,2007; Beck et al.,2009; Contreras et al.,2009; Kragl et al.,2009; Calve et al.,2010). In this study, we examined zebrafish caudal fin regeneration after repeated injuries at different ages. We demonstrated that the potential to initiate regeneration (e.g., fin outgrowth and marker gene expression) did not decline in fish that received multiple amputations. Also, the initial process of regeneration did not seem to correlate with age. However, slight differences in fin outgrowth were seen during late regeneration stages between young and old animals. At 30 dpa, for example, the regenerating caudal fin grew to approximately 95–98% of its original length in young animals (4 months old), whereas outgrowth of the caudal fin was only 88% of the original length in old animals (18 months old).
The mechanisms that underlie unlimited caudal fin regeneration in adult zebrafish remain to be further studied. We suspect that both genetic and epigenetic cues may play a role (see Akimenko et al.,1995,2003; Makino et al.,2005; Bouzaffour et al.,2009). It is possible that fin regeneration (at least the initial events) is regulated by activation of a genetic program, which may result in unlimited production of blastema cells. Through a mutant screening, for example, Whitehead et al. (2005) identified a zebrafish mutation (devoid of blastema, which is a null allele of fgf20) that blocks fin regeneration. They demonstrated that up-regulation of fgf20 is required to initiate the process of regeneration (i.e., epithelialization and blastema formation). During blastema formation, fgf20a co-expresses with msxb. Increase of msxb expression is required for tissue outgrowth during regeneration (Thummel et al.,2010). Genetic control of organ regeneration has been reported in other zebrafish organs, such as heart, spinal cord, and retina (Curado et al.,2007; Schoenebeck et al.,2007; Qin et al.,2009; Jopling et al.,2010; Montgomery et al.,2010).
In most vertebrate species, the potential to regenerate injured tissues decreases with age, which is likely due to reduced cell proliferation and differentiation (Iakova et al.,2003; Rando,2006). In zebrafish, however, the process of regenerating injured caudal fins does not seem to attenuate with age (see also Kishi et al.,2003; Keller and Murtha,2004; Lund et al.,2009). This may be due to weak but chronic expression of genes involved in tissue regeneration. We observed msxb and fgf20a expression in uninjured tissues in both young and old animals (by RT-PCR analysis; data not shown). It has been demonstrated that msx- and fgf-related gene transcripts co-express in undifferentiated mitotic cells (Nechiporuk et al.,2003; Whitehead et al.,2005). Therefore, upon injury, the expression of msxb and fgf20a is readily up-regulated to induce fin regeneration.
In summary, by using morphological and gene expression analyses, we provide evidence for unlimited fin regeneration in adult zebrafish. Unlike mammalian species, the zebrafish is capable of regenerating its tissues after repeated injuries.
Fish Care and Fin Amputation
AB zebrafish (Danio rerio) were used in this study. The fish were kept in re-circulating fish water (distilled water with Instant Ocean salt) and were fed twice per day with freshly hatched brine shrimp. Prior to fin amputations, the fish were anesthetized in 0.1% tricaine and then placed on a wet towel. Using a razor blade, approximately 50% of the caudal fin was amputated. The fish were allowed to recover in tanks placed in an incubator set at 33°C (Johnson and Weston,1995). Fin regeneration (tissue outgrowth and the expression of regeneration marker genes) was examined at different times post-amputation and the statistical differences (between different age groups or at different post-amputation times) were determined by the Student's t-test.
In Situ Hybridization
Whole-mount in situ hybridization was performed as described previously (Li et al.,2008). We used a 311-bp msxb cDNA fragment (Forward 5′-CGAGGAGGAGATGTGAA AGAT-3, Reverse 5′-GAGGAAAGTCAAAAGCC GA-3′) and a 575-bp fgf20a cDNA fragment (Forward 5′-CAGCTTCTCT CACGGCTTGG-3′, Reverse 5′-AAAG CTCAG GAACTCGCTCTG-3′) to generate anti-sense RNA probes that hybridize to msxb and fgf20a, respectively. In situ hybridization was performed in control tissues (freshly isolated caudal fins from uninjured fish, designated 0 hpa) and regenerating tissues that were collected at 24, 48, 72, and 96 hpa, respectively.
Total RNA was extracted from caudal fins (n = 4 for each PCR) using Trizol according to the manufacturer's protocol (Invitrogen, Carlsbad, CA), and was reverse-transcribed by M-MLV reverse transcriptase (Promega, Madison, WI) using the oligo (dT) primers. Quantitative PCR was performed using the SYBR Green Labeling System (BioRad, Hercules, CA). RT-PCR conditions included a denaturing step at 94°C for 2 min, 40 cycles of 94°C for 30 sec, 62°C for 30 sec, and 72°C for 30 sec for real time plate read, and a final extension at 72°C for 5 min. β-actin was used for data normalization. Primer sequences included: msxb, Forward 5′-GACGACAGTGAA GAACTAAGCG-3′, Reverse 5′-CCGTT CGGCGATAGAG AGGT-3′; fgf20a, Forward 5′-AGGAAGGACCACAGCA GATTTG-3′, Reverse 5′-CATGC CGA TACAGGTTAGAAGAGT-3′; bmp2b, Forward 5′-TCTCACGGTGCTGTTGC TCG-3′, Reverse 5′-GATTTGCTTGGG GTGGGTTT-3′; β-actin, Forward 5′-TTCACCAC CACAGCCGAAAGA-3′, Reverse 5′-TACCGCAAGATTCCATA CCCA-3′.
The authors thank Ms. Aprell Carr and Ms. April DeLaPaz for proofreading the manuscript.