Zebrafish fin regeneration involves generic and regeneration-specific osteoblast injury responses

  1. Ivonne Sehring  Is a corresponding author
  2. Hossein Falah Mohammadi
  3. Melanie Haffner-Luntzer
  4. Anita Ignatius
  5. Markus Huber-Lang
  6. Gilbert Weidinger  Is a corresponding author
  1. Institute of Biochemistry and Molecular Biology, University of Ulm, Germany
  2. Institute of Orthopaedic Research and Biomechanics, University Hospital Ulm, Germany
  3. Institute of Clinical and Experimental Trauma-Immunology (ITI), University Hospital Ulm, Germany

Abstract

Successful regeneration requires the coordinated execution of multiple cellular responses to injury. In amputated zebrafish fins, mature osteoblasts dedifferentiate, migrate towards the injury, and form proliferative osteogenic blastema cells. We show that osteoblast migration is preceded by cell elongation and alignment along the proximodistal axis, which require actomyosin, but not microtubule (MT) turnover. Surprisingly, osteoblast dedifferentiation and migration can be uncoupled. Using pharmacological and genetic interventions, we found that NF-ĸB and retinoic acid signalling regulate dedifferentiation without affecting migration, while the complement system and actomyosin dynamics affect migration but not dedifferentiation. Furthermore, by removing bone at two locations within a fin ray, we established an injury model containing two injury sites. We found that osteoblasts dedifferentiate at and migrate towards both sites, while accumulation of osteogenic progenitor cells and regenerative bone formation only occur at the distal-facing injury. Together, these data indicate that osteoblast dedifferentiation and migration represent generic injury responses that are differentially regulated and can occur independently of each other and of regenerative growth. We conclude that successful fin bone regeneration appears to involve the coordinated execution of generic and regeneration-specific responses of osteoblasts to injury.

Editor's evaluation

This work is of interest to readers in the field of bone regeneration, and more broadly to readers in the field of tissue repair and regenerative medicine. The authors took advantage of a well–established in vivo model, live imaging, pharmacological inhibition, and genetic strategies to dissect the interrelations of key cellular events in zebrafish fin regeneration. The finding of how distinct generic injury responses are differentially regulated and are functioning independently from each other, is a valuable piece of information for the community.

https://doi.org/10.7554/eLife.77614.sa0

Introduction

For humans and other mammals, the traumatic loss of a limb represents an irreversible and permanent defect, as lost appendages cannot be restored. In contrast, teleost fish and urodele amphibians are able to fully regenerate limbs/fins. Thus, the zebrafish caudal fin has become a popular model to study the restoration of bone tissue (Gemberling et al., 2013; Pfefferli and Jaźwińska, 2015). The fin skeleton consists of dermal (directly ossifying) skeletal elements, the fin rays or lepidotrichia, which make up the largest part of the fin, and endochondral parts close to the body. Rays are segmented with flexible joints, and each segment consists of two concave hemirays which are lined by a single layer of osteoblasts on the inner and outer surface (Figure 1A). Within 1 day after fin amputation, a wound epidermis covers the injured tissue. Next, a blastema forms atop of each ray, which contains the proliferative source cells for regeneration of the ray skeleton. Within hours after amputation, osteoblasts close to the amputation plane dedifferentiate, that is, they downregulate the expression of the mature osteoblast marker bglap, upregulate preosteoblast markers like runx2, and become proliferative (Knopf et al., 2011; Sousa et al., 2011; Stewart and Stankunas, 2012). Furthermore, osteoblasts relocate towards the amputation plane and beyond to contribute to the blastema (Knopf et al., 2011; Geurtzen et al., 2014). In the regenerate, these dedifferentiated osteoblasts remain lineage-restricted and redifferentiate to osteoblasts (Knopf et al., 2011; Stewart and Stankunas, 2012). While both osteoblast dedifferentiation and migration occur in response to fin amputation and bone fractures in zebrafish (Knopf et al., 2011; Geurtzen et al., 2014), the interrelation of these processes is not understood; specifically, whether dedifferentiation is a requirement for migration is not known.

Figure 1 with 1 supplement see all
Osteoblasts dedifferentiate and migrate in response to fin amputation.

(A) Schematic presentation of one fin ray with mature osteoblasts expressing bglap and entpd5 lining the hemirays. (B) RNAscope in situ analysis of bglap expression in segments 0, –1, and –2 at 1 day post amputation (dpa). Expression of bglap gradually decreases towards the amputation plane. N (experiments)=1, n (fins)=5, n (rays)=15. Error bars represent SEM. Scale bar, 100 µm. (C) RNAscope in situ detection of entpd5 expression in segments 0, –1, and –2 at 1 dpa. Scale bar, 100 µm. (D) In bglap:GFP fish, expression of gfp RNA is downregulated in segment –1 at 2 dpa, while GFP protein persists. Segment –2 shows overlap of gfp RNA and GFP protein (note that GFP protein is enriched in the nuclei). Dashed lines indicate segment border. Distal to the right. Scale bar, 50 µm (E) Scheme illustrating the use of bglap:GFP transgenics for analysis of osteoblast migration via relocation of cells that are negative for gfp RNA, but positive for GFP protein. (F) At 0 dpa, entpd5:Kaede positive osteoblasts in the proximal half of segment –1 were photoconverted from entpd5:kaedeGreen to entpd5:kaedeRed. Repeated imaging reveals distal relocation of photoconverted osteoblasts by 1 dpa. Dashed lines indicate segment borders. Distal to the right. Scale bar, 50 µm. (G) bglap:GFP is expressed in a subset of osteoblasts in the centre of each segment and absent around the joints. Repeated imaging reveals relocation of GFP+ cells towards the distal joint in segment –1 by 1 dpa. Distal to the right. Scale bar, 10 µm. (H) Quantification of the distance between the bulk of bglap:GFP+ cells and joints in segment –1 at 0 and 1 dpa. At 1 dpa, the proximal distance is increased, while the distal distance is reduced. N (experiments)=3, n (fins)=26, n (rays)=26. Error bars represent 95% CI. Unpaired t-test. (I) Quantification of bglap:GFP+ bulk migration in segment −3, –2, and –1 at 1 dpa. 100% indicates full crossing of the distance to the respective joint. Osteoblasts migrate distally in segment –1. N (experiments)=2, n (fins)=19, n (rays)=29. Error bars represent 95% CI. Mann Whitney test.

Cell migration requires formation and retraction of membrane protrusions, which are regulated by the dynamic actomyosin network, while microtubuli play a role in organising the polarisation of migrating cells (Petrie et al., 2009). Bone tissue is permanently turned over by bone remodelling, a process of alternating bone resorption and bone formation (Kular et al., 2012). For bone formation, osteoblast precursors migrate to the resorbed sites (Dirckx et al., 2013). Similarly, during mammalian fracture healing, osteoblasts are recruited to the site of injury (Thiel et al., 2018). Several factors have been shown to act as chemoattractants for osteoblasts in vitro, but few have been confirmed to play a role in vivo (Dirckx et al., 2013; Thiel et al., 2018). One candidate osteoblast guidance cue is the complement system, which represents the major fluid phase part of innate immunity and is activated immediately after injury. It consists of more than 50 proteins, including serial proteases, whose activation leads to the formation of the peptides C3a and C5a, generated from the precursors C3 and C5, respectively (Thorgersen et al., 2019). While the majority of C3 and C5 precursors are expressed in the liver and distributed to the periphery via the circulation (Merle et al., 2015), C3 and C5 mRNA are also expressed by human osteoblasts, which in addition can cleave native C5 into C5a (Ignatius et al., 2011b). During bone fracture healing in mammals, the receptor for C5a (C5aR) is expressed by osteoblasts, and C5a can act as chemoattractant for osteoblasts in vitro (Ignatius et al., 2011b). In C5-deficient mice, bone repair after fracture is severely impaired (Ehrnthaller et al., 2013); however, whether this is due to defects in osteoblast migration is not yet known.

In this study, we analysed injury-induced osteoblast migration in vivo in the regenerating zebrafish fin. We show that osteoblast cell shape changes and migration depend on a dynamic actomyosin cytoskeleton, but not on microtubuli turnover. Pharmacological interference with C3a and C5a suggests that the complement system regulates osteoblast migration in vivo. Using genetic and pharmacological manipulation of NF-κB, retinoic acid (RA), and complement signalling, we found that dedifferentiation and migration can be uncoupled and are independently regulated, suggesting that dedifferentiation is not a prerequisite for migration. Furthermore, we established a novel injury model in which an internal bone defect within fin rays allows us to study osteoblast behaviours at proximally and distally facing injuries. Intriguingly, osteoblast migration and dedifferentiation occur at both injury sites, yet only at the distal injury a preosteoblast population forms and only here regenerative growth commences. We conclude that osteoblast migration and dedifferentiation represent generic injury responses in zebrafish, and that successful bone regeneration depends on additional, regeneration-specific events.

Results

Osteoblasts elongate, align along the proximodistal axis, and migrate towards the amputation plane

In non-injured caudal fins, differentiated osteoblasts expressing the markers bglap and entpd5 line the two segmented hemirays that together form the fin rays (Figure 1A). We use the following terminology to describe the subsequent experiments: segment 0 is the fin ray segment through which we amputate (at 50% of its length), and segments −1, –2, and –3 are located further proximally (Figure 1A). Note that only segment 0 is mechanically affected by the amputation injury. We have previously shown that osteoblasts dedifferentiate in response to fin amputation, that is they revert from a mature, non-proliferative state into an undifferentiated progenitor-like state, which includes loss of bglap expression and upregulation of the preosteoblast marker runx2 (Knopf et al., 2011; Geurtzen et al., 2014). Using RNAscope in situ hybridisation, we can now show that downregulation of bglap occurs in a graded manner and that entpd5 expression is similarly downregulated during dedifferentiation (Figure 1B and C). At 1 day postamputation (1 dpa), expression of entpd5 and bglap remains high in segment –2, but gradually decreases towards the amputation plane and is almost entirely absent from segment 0, with entpd5 downregulation being more pronounced (Figure 1B and C). While RNA expression of these genes is downregulated within hours after injury, GFP or Kaede fluorescent proteins (FPs) expressed in bglap or entpd5 reporter transgenic lines persist for up to 3 days, even though transgene transcription is shut down rapidly as well (Knopf et al., 2011). We can confirm these earlier findings using the more sensitive RNAscope in situs. In bglap:GFP transgenics at 2 dpa, gfp RNA and GFP protein colocalised to the same cells in segment –2, where osteoblasts do not dedifferentiate (Figure 1D). In contrast, in the distal segment –1 GFP protein was present, but barely any gfp transcript could be detected (Figure 1D). Thus, persistence of GFP in bglap:GFP transgenics can be used for short-term tracing of dedifferentiated osteoblasts in zebrafish (Figure 1E). At 1 dpa, bglap:GFP+ cells upregulated expression of the preosteoblast marker runx2a and of cyp26b1, an enzyme involved in RA signalling (Blum and Begemann, 2015), which regulates dedifferentiation (Figure 1—figure supplement 1A,B). Both markers were exclusively upregulated in segment –1 and segment 0 at 1 dpa, but were absent in segment –2. Together, these data show that osteoblasts in segment –1 and segment 0 lose expression of mature markers and gain expression of dedifferentiation markers.

We have previously provided evidence that osteoblasts close to the amputation plane do not only dedifferentiate, but also relocate towards the injury site after fin amputation. To trace osteoblasts, we used the transgenic line entpd5:kaede (Geurtzen et al., 2014), in which Kaede fluorescence can be converted from green to red by UV light (Ando et al., 2002). We photoconverted osteoblasts in the proximal half of segment –1, while osteoblasts in the distal half remained green (Figure 1F). At 1 dpa, red osteoblasts were found in the distal half (Figure 1F), showing that photoconverted osteoblasts had relocated distally. To quantify osteoblast relocation, we used the bglap:GFP transgenic line (Knopf et al., 2011). In adult non-injured fin rays, bglap:GFP is expressed in a subset of mature osteoblasts in the centre of every ray segment (Figure 1G). The restriction of bglap:GFP+ cells to the segment centre results in a zone devoid of bglap:GFP+ cells at both ends of a segment, and its invasion by GFP+ cells can be used as a read-out for bulk migration of osteoblasts after fin injury. Within 1 dpa, the distance between the GFP+ cells and the distal joint was reduced in segment –1, while at the proximal joint, the distance increased (Figure 1G and H). Therefore, osteoblasts did not spread out across the segment, but were relocated directionally towards the amputation plane. We also analysed if osteoblasts in more proximal segments (further away from the amputation plane) move distally and quantified their relocation. 100% relocation indicates that the distal front of the GFP+ bulk of cells has reached the respective distal joint. At 1 dpa, osteoblasts in segment –1 relocated distally, while no movement could be detected in segments –3 and –2 (Figure 1I). In summary, osteoblasts respond to fin amputation by dedifferentiation (detected by downregulation of transcription of differentiation markers, as well as upregulation of preosteoblast markers) and by movement towards the amputation plane, as detected by relocation of GFP+ cells (Figure 1E). Dedifferentiation and migration occur in a region encompassing about 350 µm proximal to the amputation plane, comprising the amputated segment 0 and the adjacent segment –1.

To observe osteoblast behaviour at single cell resolution in live fish after fin amputation, we generated double transgenic fish expressing bglap:GFP (Knopf et al., 2011) and entpd5:kaede in mature osteoblasts. Expression of two fluorescent proteins increased signal for repeated imaging, and partial photoconversion of kaedeGreen into kaedeRed resulted in a different colouring for each osteoblast, which facilitated tracking of morphology changes at single cell resolution. Live imaging revealed the formation of long protrusions, lasting for at least 2 hr and extending towards the amputation site, and the directed movement of cell bodies relative to their surroundings (Video 1, Figure 2—figure supplement 1A). We conclude that osteoblasts relocate by active migration. Migrating cells typically possess an identifiable cell front and a cell rear along an axis approximately aligned with the direction of locomotion. We found that bglap:GFP+ osteoblasts changed their shape after amputation. In a mature, non-injured segment, osteoblasts were roundish, and they retained this morphology after amputation in segment –3 and segment –2 at 1 dpa, as determined by a width/length ratio of ~0.4 (Figure 2A and B, Figure 2—figure supplement 1B). In contrast, at 1 dpa osteoblasts in segment –1 displayed an elongated shape (width/length ratio ~0.2) and they presented long extensions (Figure 2A and B). The elongation of osteoblasts occurred in alignment with the proximodistal axis of the fin, as evident by an angle reduction between this axis and the long axis of the osteoblasts (Figure 2C). Orientation of osteoblasts along the proximodistal axis could first be detected at 12 hr postamputation (hpa) (Figure 2—figure supplement 1C), while elongation was first observed at 15 hpa (Figure 2—figure supplement 1D). We interpret the elongation and orientation along the proximodistal axis and the formation of long-lived protrusions along this axis as events that prime osteoblasts for directed active migration towards the injury.

Video 1
Osteoblast migration.

Live imaging of osteoblasts at 1 day post amputation (dpa) in segment –1 (distal to the right) in a double transgenic line expressing bglap:GFP and entpd5:kaede. Partial conversion of kaedeGreen into kaedeRed results in a different colouring for each cell. Stationary yellow arrowheads highlight the migration of cell bodies towards the amputation plane and retraction of the rear ends. Additionally, several cells can be observed forming long protrusions extending distally (green arrowheads). Distal to the right. Scale bar, 10 µm.

Figure 2 with 2 supplements see all
Osteoblasts elongate and orient along the proximodistal axis of the fin in response to fin amputation.

(A) bglap:GFP+ osteoblast morphology in a non-injured fin (upper panel) and in segment –1 at 1 day post amputation (dpa) (lower panel). Scale bar, 10 µm. (B) Quantification of bglap:GFP+ osteoblast roundness as width/length ratio at 1 dpa. Osteoblasts are more elongated in segment –1. N (experiments)=1, n (fins)=5, n (rays)=5, n (cells)=72. Error bars represent 95% CI. Kruskal-Wallis test. (C) Quantification of bglap:GFP+ osteoblast orientation as angular deviation from the proximodistal axis at 1 dpa. In segment –1, osteoblasts display increased alignment along the axis. N (experiments)=1, n (fins)=5, n (rays)=5, n (cells)=44. Error bars represent 95% CI. Kruskal-Wallis test. (D) At 2 dpa, bglap:GFP+ osteoblasts can be observed within the distal joint of segment –1, and GFP+ protrusions can be seen spanning the joint, indicating that osteoblasts migrate through the joint. Yellow arrowheads, joints; red arrowheads, amputation plane. Scale bar, 10 µm. (E) Osteoblast roundness in distal and proximal parts of segments (40% of total segment length from the respective end) at 24 hpa. No difference within segment –1 can be detected. N (experiments)=1, n (fins)=5, n (rays)=5, n (cells)=34 (segment –2), 55 (proximal segment –1), 78 (distal segment –1). Error bars represent 95% CI. Mann-Whitney test. (F) Osteoblast roundness in distal and proximal parts of segment –1 at 15 hpa. No difference within segment –1 can be detected. N (experiments)=1, n (fins)=6, n (rays)=15, n (cells)=82 (proximal), 49 (distal). Error bars represent 95% CI. Mann-Whitney test. The observed relative difference is 0.3%, the calculated smallest significant difference 13%, which is smaller than what we observe between segment –2 and segment –1 (B, 54%).

Fins grow by the addition of new segments distally, and in the distal-most, youngest segment of non-injured fins, bglap:GFP is not expressed (Knopf et al., 2011). Similarly, in the regenerating fins increasing numbers of GFP+ cells can be detected in more proximally located, older segments, reflecting the progressive differentiation of osteoblasts with time (Figure 2—figure supplement 2A). In less mature segments, the pre-osteoblast marker Runx2 can be detected, but its expression does not overlap with bglap:GFP expression in mature osteoblasts (Figure 2—figure supplement 2B). Importantly, in newly regenerated segments that start to upregulate bglap expression, all bglap:GFP+ cells appear in the centre of the segments; we could not observe GFP+ cells within joints (Figure 2—figure supplement 2A). This suggests that bglap:GFP+ osteoblasts from older segments do not migrate into less mature segments during formation of new segments in the course of fin growth. Rather, the mature osteoblast population in a segment arises via differentiation of osteoblasts at the position within the segment at which they were formed during segment addition. In contrast, within 2 days after fin amputation, bglap:GFP+ cells appeared in the fin stump within the joint between segment –1 and segment 0 (Figure 2D, yellow arrowhead), indicating that GFP+ osteoblasts from segment –1 crossed the joint during their migration towards the amputation plane. Thus, migration of mature osteoblasts observed after fin amputation or bone fracture (Geurtzen et al., 2014) appears to represent a specific early response to injury, while addition of new bony segments at the distal tip of growing fins during ontogeny or regeneration does not involve migration of differentiated osteoblasts.

Since we found that differentiation markers are gradually downregulated towards the amputation plane at 1 dpa (Figure 1B and C), we wondered whether a similar gradient can be observed for cell morphology changes. However, osteoblasts were elongated to the same extent at the proximal and distal end of segment –1 at 24 hpa (Figure 2E). As elongation was first observed at 15 hpa (Figure 2—figure supplement 1D), we also analysed osteoblasts in proximal and distal regions of segment –1 at this time point. Yet, no morphological differences in osteoblasts of proximal and distal regions of segment –1 were detected at 15 hpa (Figure 2G). We conclude that dedifferentiation and cell shape changes are responses of osteoblasts to injury that occur in the same cells, yet the magnitude of these responses can vary. While cell shape change is a binary response that affects all osteoblasts in the responsive zone equally, dedifferentiation is a graded process.

Actomyosin, but not microtubule dynamics is required for osteoblast cell shape changes and migration

One of the primary forces facilitating cell shape changes and cell motility is the myosin-associated actin cytoskeleton (Chugh and Paluch, 2018). To interfere with the treadmilling of actin microfilaments (F-actin), we used cytochalasin D, which binds to F-actin and disturbs its turnover (Brown and Spudich, 1979; Brown and Spudich, 1981). Drug treatment strongly impaired osteoblast elongation in segment –1 at 1 dpa, while it did not affect osteoblast morphology in segment –2 and segment –3 (Figure 3A). Concomitantly, treatment with cytochalasin D also resulted in reduced alignment of osteoblasts along the proximodistal axis in segment –1, but did not affect the orientation of osteoblasts in segment –2 and segment –3 (Figure 3B). Interference with actin dynamics also significantly reduced the bulk migration of bglap:GFP+ cells in segment –1 (Figure 3C). To interfere with another hallmark of a dynamic cytoskeleton, the contractility of the actomyosin network, we injected fish with blebbistatin, an inhibitor of myosin II ATPase (Kovács et al., 2004). The drug did not affect osteoblast cell shape in segments –3 and –2, but impaired the elongation and reorientation of osteoblasts along the proximodistal axis in segment –1 (Figure 3D and E). Concomitantly, bulk osteoblast migration was reduced (Figure 3C).

Figure 3 with 1 supplement see all
Interference with actomyosin, but not microtubule dynamics inhibits osteoblast cell shape changes and migration.

(A) Osteoblast roundness at 1 day post amputation (dpa). Inhibition of actin dynamics with cytochalasin D does not alter osteoblast cell shape in segments –3 and –2, but cell elongation in segment –1 is inhibited. N (experiments)=3, n (fins)=15, n (rays)=15, n (cells)=87. Error bars represent 95% CI. Kruskal-Wallis test. (B) Osteoblast orientation at 1 dpa. Cytochalasin D does not alter osteoblast orientation in segments –3 and –2, but alignment along the proximodistal axis in segment –1 is impaired. N (experiments)=3, n (fins)=15, n (rays)=15, n (cells)=93. Error bars represent 95% CI. Kruskal-Wallis test. (C) Both cytochalasin D and blebbistatin treatment impair bulk osteoblast migration. Images show overlay of 0 dpa (green) and 1 dpa (pink) pictures, with the pink arrowhead indicating relocation of osteoblasts in controls, where no signal overlap is observed at the distal side. Graph: 100% indicates full crossing of the distance to the respective joint at 1 dpa. Cytochalasin D: N(experiments)=3, n (fins)=22, n (rays)=44; blebbistatin: n (fins)=24, n (rays)=48, appertaining controls have the same n. Error bars represent 95% CI. Mann-Whitney test. Scale bar, 100 µm. (D) Osteoblast roundness at 1 dpa. Inhibition of myosin activity with blebbistatin does not alter osteoblast cell shape in segments –3 and –2, but cell elongation in segment –1 is inhibited. N (experiments)=3, n (fins)=15, n (rays)=15, n (cells)=89. Error bars represent 95% CI. Kruskal-Wallis test. (E) Osteoblast orientation at 1 dpa. Blebbistatin does not alter osteoblast orientation in segments –3 and –2, but alignment along the proximodistal axis in segment –1 is impaired. N (experiments)=3, n (fins)=15, n (rays)=15, n (cells)=77. Error bars represent 95% CI. Kruskal-Wallis test. (F) Neither cytochalasin nor blebbistatin treatment affects osteoblast dedifferentiation. Plotted are the 1 dpa bglap RNAscope signal intensity levels in segment –1 relative to those in the same ray in segment –2. N (experiments)=1, n (fins)=6, n (rays)=12. Error bars represent 95% CI. Kruskal-Wallis test. The observed relative difference is 3% (blebbistatin) and 2% (cytochalasin), the calculated smallest significant difference for both assays is 9%, which is smaller than what we observe after retinoic acid treatment (Figure 4B, 63%). (G) Inhibition of microtubule dynamics with nocodazole does not affect osteoblast migration. Images show overlay of 0 dpa (green) and 1 dpa (pink) pictures. N (experiments)=1, n (fins)=10, n (rays)=20. Error bars represent 95% CI. Unpaired t-test. The observed relative difference is 1%, the calculated smallest significant difference is 24% and thus smaller than what we observe in actomyosin treatment regimens (C, 60–63%). Scale bar, 100 µm.

Due to osteoblast dedifferentiation, at 1 dpa bglap RNA expression in segment –1 is reduced to ~50% compared to the expression in segment –2 (Figure 3—figure supplement 1A). Interestingly, downregulation of bglap expression was not affected by either cytochalasin D or blebbistatin treatment (Figure 3F), indicating that their effect on cell shape change and migration is not secondary to impaired dedifferentiation. Likewise, upregulation of Runx2 at 2 dpa was not diminished upon cytochalasin D treatment (Figure 3—figure supplement 1B). Yet, regenerative growth was reduced at 3 dpa (Figure 3—figure supplement 1C).

Besides the actomyosin network, microtubules (MT) are an important force to drive cell shape changes and cell motility (Etienne-Manneville, 2013). To analyse a potential role of MT in the amputation-induced migration of osteoblasts, we treated fish with nocodazole, a drug that disrupts MT assembly/disassembly (Florian and Mitchison, 2016), using a regime which was shown to be effective in zebrafish (Poss et al., 2004). At 1 dpa, we could not detect a difference in the extent of migration in drug-treated fish compared to controls (Figure 3G). Together, these data indicate that actomyosin, but not microtubule dynamics is required for injury-induced osteoblast migration.

Osteoblast migration is independent of osteoblast dedifferentiation

We next wondered whether dedifferentiation is a prerequisite for osteoblasts to change their shape and to migrate. We have previously shown that NF-κB signalling negatively regulates osteoblast dedifferentiation (Mishra et al., 2020). We induced expression of an inhibitor of NF-κB signalling (IkB super repressor [IkBSR]); (Van Antwerp et al., 1996) specifically in osteoblasts, using the tamoxifen-inducible Cre line OlSp7:CreERT2-p2a-mCherrytud8 (osx:CreER) (Knopf et al., 2011) and an ubiquitously expressed responder line hsp70l:loxP Luc2-myc Stop loxP nYPet-p2a-IκBSR, cryaa:AmCyanulm15Tg, which expresses IkBSR and nYPet after recombination (hs:Luc to nYPet IκBSR). NF-kB signalling activation was induced by expressing constitutively active human IKK2 (IkB kinase) and nuclear localised BFP after recombination in osteoblasts using hsp70l:loxP Luc-myc Stop loxP IKKca-t2a-nls-mTagBFP2-V5, cryaa:AmCyanulm12Tg (hs:Luc to IKKca BFP) fish (Mishra et al., 2020). Expression of GFP driven by the Cre-responder line hsp70l:loxP DsRed2 loxP nlsEGFPtud9 (hs:R to G) (Knopf et al., 2011) served as a negative control. Using these tools, we have previously shown that activation of NF-κB-signalling in osteoblasts inhibits their dedifferentiation, while its suppression promotes osteoblast dedifferentiation (Mishra et al., 2020). We used the same setup to ask whether NF-κB signalling also regulates osteoblast migration. Mosaic recombination allowed us to compare recombined and non-recombined cells within the same segment. Analysis of cell shape in the control line hs:R to G revealed elongation of both non-recombined and recombined cells in segment –1 compared to segment –2 (Figure 4A). Neither pathway inhibition by expression of IkBSR nor forced activation by expression of IKKca affected osteoblast elongation in segment –1. We conclude that NF-κB signalling regulates osteoblast dedifferentiation, but not the cell shape changes associated with osteoblast migration.

NF-κB and retinoic acid signalling regulate osteoblast dedifferentiation but not migration.

(A) Mosaic recombination in osx:CreER; hs:R to G, osx:CreER; hs:Luc to IKKca BFP and osx:CreER; hs:Luc to nYPet IκBSR fish. Zns5 labels the membrane of osteoblasts. White arrowheads highlight roundish osteoblasts, white arrows elongated osteoblasts. Osteoblast roundness in recombined and non-recombined osteoblasts in segment –2 and segment –1 at 1 day post amputation (dpa) is plotted. Recombined osteoblasts expressing IKKca (marked by BFP) or IκBSR (marked by nYPet) elongate in segment –1 to a similar extent as osteoblasts expressing the negative control GFP. N (experiments)=1, R to G: n (fins)=3, n (rays)=10, n (cells)=406; IKKca: n (fins)=4, n (rays)=7, n (cells)=244; IκBSR: n (fins)=8, n (rays)=18, n (cells)=246. Error bars represent 95% CI. Kruskal-Wallis test. Scale bar, 10 µm. (B) Treatment with retinoic acid (RA) inhibits osteoblast dedifferentiation measured as bglap RNAscope intensity at 1 dpa in segment –1 relative to the intensity in segment –2 in the same rays. N (experiments)=1, n (fins)=6, n (rays)=12. Error bars represent 95% CI. Unpaired t-test. (C) Osteoblast roundness at 1 dpa. RA treatment does neither alter osteoblast cell shape in segments –3 and –2, nor elongation in segment –1. N (experiments)=2, n (fins)=10, n (rays)=10, n (cells)=83. Error bars represent 95% CI. Kruskal-Wallis test. The observed relative difference in segment –1 is 2%, the calculated smallest significant difference is 7%, which is smaller than what we observe in cytochalasin treatment regimens (Figure 3A, 22%). (D) Osteoblast orientation at 1 dpa. RA treatment does not affect osteoblast orientation in segments –3 and –2, nor alignment along the proximodistal axis in segment –1. N (experiments)=2, n (fins)=10, n (rays)=10, n (cells)=89. Error bars represent 95% CI. Kruskal-Wallis test. The observed relative difference in segment –1 is 1%, the calculated smallest significant difference is 7%, which is smaller than what we observe in cytochalasin treatment regimens (Figure 3B, 32%). (E) RA treatment does not affect bulk migration of osteoblasts towards the amputation plane in segment –1. N (experiments)=3, n (fins)=26, n (rays)=52. Error bars represent 95% CI. Unpaired t-test. The observed relative difference is 2%, the calculated smallest significant difference is 22%, which is smaller than what we observe in actomyosin treatment regimens (Figure 3C 60–63%).

RA signalling inhibits osteoblast dedifferentiation downstream of NF-κB signalling (Blum and Begemann, 2015; Mishra et al., 2020). Treatment with RA impaired osteoblast dedifferentiation after amputation, as bglap RNA remained at higher levels in segment –1 (Figure 4B). In contrast, osteoblast elongation and alignment along the proximodistal axis in segment –1 at 1 dpa were not affected (Figure 4C and D). Similarly, RA did not affect bulk osteoblast migration towards the amputation plane (Figure 4E).

Together, the data on manipulation of actomyosin dynamics, NF-κB, and RA signalling suggest that osteoblast dedifferentiation and migration are independently regulated, as one osteoblast injury response can be impaired without affecting the other. Furthermore, they indicate that osteoblast dedifferentiation is not a prerequisite for migration.

The complement system is required for osteoblast migration in vivo

As shown above, we found that osteoblast cell shape changes and migration are directed towards the amputation site. Directional cell migration is usually initiated in response to extracellular cues such as chemokines or signals from the extracellular matrix (Swaney et al., 2010). As part of the innate immune system, the complement system is activated immediately after injury, and activated complement factors can act as chemoattractants for osteoblasts in vitro (Ignatius et al., 2011a). We thus asked whether the complement system regulates osteoblast migration in vivo. The zebrafish genome contains homologues of all fundamental mammalian complement components (Boshra et al., 2006; Zhang and Cui, 2014), including the central components c3 and c5 and the corresponding receptors c3aR and c5aR1, respectively. A second c5a receptor, c5aR2, has so far only been characterised in mammals, and sequence database interrogation indicates that it is absent in zebrafish. RNAscope in situ analysis revealed that c5aR1 is expressed in mature osteoblasts (Figure 5A), as well as in other cells of the fin (Figure 5—figure supplement 1A). Similarly, quantitative RT-PCR (qRT-PCR) on GFP+ and GFP− cells sorted from 1 dpa bglap:GFP fins revealed that c5aR1 is expressed in both fractions, and thus in osteoblasts and other cell types (Figure 5—figure supplement 1B). Likewise, expression of c3aR1 could be detected in both fractions (Figure 5—figure supplement 1B). As measured by qRT-PCR, transcription of the complement factor precursor c5 and the six c3a precursor paralogues could readily be detected in the liver, but was minimal in non-injured fins (103–106-fold less than in the liver, Figure 5B). Yet, we could observe an upregulation of c5 and c3a.5 expression in the fin at 6 hpa (Figure 5B). Thus, while the majority of complement components that are available for injury-induced activation in the fin are probably produced in the liver and delivered to the fin by the circulation, injury-induced local expression in the fin might contribute to activation of the system as well.

Figure 5 with 4 supplements see all
Complement system signalling regulates osteoblast cell shape changes and migration in vivo.

(A) RNAscope in situ detection of c5aR1 expression in bglap expressing osteoblasts in a non-injured fin segment. YZ section shows localisation of both RNAs in the same cell layer. Scale bar, 10 µm. (B) Expression of complement factors detected by qRT-PCR in the liver, fins at 0 hpa and 6 hpa. Plotted are the ΔΔCT values relative to the 0 hpa fins. Each data point represents one biological replicate. Error bars represent SD. Two-way ANOVA. (C) Osteoblast roundness at 1 day post amputation (dpa). The C5aR1 inhibitor W54011 does not alter osteoblast cell shape in segments –3 and –2, but cell elongation in segment –1 is inhibited. N (experiments)=3, n (fins)=5, n (rays)=5, n (cells)=116. Error bars represent 95% CI. Kruskal-Wallis test. (D) Osteoblast roundness at 1 dpa. The C3R inhibitor SB290157 does not alter osteoblast cell shape in segments –3 and –2, but cell elongation in segment –1 is inhibited. N (experiments)=1, n (fins)=5, n (rays)=5, n (cells)=38. Error bars represent 95% CI. Kruskal-Wallis test. (E) Inhibition of C5aR1 with either W54011 or PMX205, and inhibition of C3aR with SB290157 impairs bulk osteoblast migration. Images show overlay of 0 dpa (green) and 1 dpa (pink) pictures. N (experiments)=3, W54011: n (fins)=27, n (rays)=54; PMX-205, SB290157: n (fins)=29, n (rays)=58. Error bars represent 95% CI. Mann-Whitney tests. Scale bar, 100 µm. (F) Neither inhibition of C5aR1 (W54011, PMX205) nor of C3aR (SB290157) affects osteoblast dedifferentiation measured as bglap RNAscope intensity at 1 dpa in segment –1 relative to the intensity in segment –2 of the same rays. N (experiments)=1; n (fins)=5 (DMSO), 4 (PMX205), 6 (SB290157); n (rays)=12. Error bars represent 95% CI. Kruskal-Wallis test. The observed relative difference is 1% for PMX205 and SB290157, the calculated smallest significant differences are 6% (PMX205) and 5% (SB290157), which are smaller than what we observe after retinoic acid treatment (Figure 4B; 7%).

Treatment with W54011, a specific C5aR1 antagonist (Sumichika et al., 2002), impaired osteoblast elongation in segment –1 at 1 dpa without affecting osteoblast cell shape in segments –2 and –3 (Figure 5C). As also the complement factor C3a can act as chemoattractant after injury (Huber-Lang et al., 2013), we treated fish with SB290157, a specific antagonist of the complement receptor C3aR (Ames et al., 2001). Osteoblast elongation was impaired in segment –1 at 1 dpa, while cell shape in segment –2 and segment –3 was not affected (Figure 5D). Both drugs also significantly reduced bulk osteoblast migration (Figure 5E). To confirm these findings, we repeated the migration assay with PMX205, another C5aR1 antagonist (March, 2004; Jain et al., 2013), which also reduced osteoblast migration at 1 dpa (Figure 5E). Together, these data strongly suggest that the complement system regulates injury-induced directed osteoblast migration in vivo.

Interestingly, none of the complement inhibitors affected osteoblast dedifferentiation as quantified by reduction of bglap RNA expression in segment –1 (Figure 5F). This supports the view that osteoblast dedifferentiation and migration are independent responses of osteoblasts to injury. Complement components can also contribute to the regulation of cell proliferation and tissue regeneration (Mastellos and Lambris, 2002). In the non-injured fin, bglap:GFP+ osteoblasts are non-proliferative, but upon amputation osteoblasts proliferate at 2 dpa (Figure 5—figure supplement 2A ,B). Proliferation is restricted to segment –1 and segment 0 (Figure 5—figure supplement 2C), and RNAscope in situ analysis of bglap expression revealed that the majority of EdU+ osteoblasts have strongly downregulated bglap (Figure 5—figure supplement 2D). Inhibition of C5aR1 with PMX205 had no effect on osteoblast proliferation in segment –1 at 2 dpa (Figure 5—figure supplement 3A). Furthermore, upregulation of Runx2 was not changed by PMX205 treatment (Figure 5—figure supplement 3B), and regenerative growth was not affected in fish treated with either W54011, PMX205, or SB290157 (Figure 5—figure supplement 3C). We conclude that the complement system specifically regulates injury-induced osteoblast migration, but not osteoblast dedifferentiation or proliferation in zebrafish.

Activation of the complement system is a general early response after injury that also occurs in wounds that heal, but do not trigger structural regeneration of the kind we observe after fin amputation. Thus, we wondered whether osteoblast migration can be triggered by injuries that do not induce structural regeneration. Lesions of the interray skin quickly heal (Chablais and Jazwinska, 2010; Chen et al., 2015) but do not trigger bone regeneration. Yet, such wounds can induce signals that are sufficient to trigger structural regeneration in a missing tissue context (Owlarn et al., 2017). We found that skin injuries induced close to the ray bone did not induce osteoblast migration, neither off the bone into the intraray nor on the bone towards the joints (Figure 5—figure supplement 4A). Osteoblasts also migrate towards fractures induced in the fin ray bone (Geurtzen et al., 2014). Interray injury combined with bone fracture resulted in osteoblast migration towards the fracture, but osteoblasts did not migrate away from the bone matrix into the interray skin (Figure 5—figure supplement 4B). We conclude that generic wounding-induced signals are not sufficient to attract osteoblasts in the absence of additional prerequisites, which might include the presence of a permissive substrate for migration.

An internal bone defect model separates generic from regeneration-specific osteoblast injury responses

In response to fin amputation, all osteoblast injury responses occur directed towards the amputation plane, that is, dedifferentiation is more pronounced distally, osteoblasts migrate distalwards, and the proliferative preosteoblast population forms distally of the amputation plane. We wondered how osteoblasts respond to injuries that occur proximal to their location. To test this, we established a fin ray injury model featuring internal bone defects. We removed hemiray segments at two locations within one ray, leaving a centre segment with two injury sites, one facing proximally, the other distally (Figure 6A). It was recently shown that in a similar cavity injury model a blastema forms only at the distal-facing injury, while the proximally facing site displays no regenerative growth (Cao et al., 2021). In our hemiray removal model, both injury sites are equally severed from the stump, and thus cut off from innervation and blood supply. Therefore, any potential differences in the injury responses at the proximal vs distal injury site cannot be explained by differences in innervation or blood circulation. Within 2 days post injury (dpi), blood flow through the centre segment was restored (Figure 6—figure supplement 1A, Video 2). At 3 dpi, Runx2+ preosteoblasts and Osterix+ committed osteoblasts accumulated almost exclusively in the defect beyond the distal injury, but not at the proximal injury (Figure 6B - D). Subsequently, new bone matrix formed exclusively at the distal site (Figure 6E).New bone matrix also formed at the distal side of the bone located proximal to the proximal bone defect (arrowhead in Figure 6A and data not shown), but not at the proximal injury site of the centre segment. Thus, our hemiray removal injury model reveals that accumulation of a preosteoblast population, and regeneration of bone occurs only at distal-facing wounds.

Figure 6 with 1 supplement see all
A hemiray removal injury model distinguishes generic vs regeneration-specific osteoblast injury responses.

(A) Hemiray removal scheme creating a proximal and distal facing injury on both sides of a central intact segment. (B) At 3 days post injury (dpi), Runx2+ preosteoblasts and committed osteoblasts expressing Osterix have only accumulated in the bone defect beyond the distal injury site of the centre segment, but not beyond the proximal injury site, as determined by immunofluorescence. Dashed line outlines centre segment. Scale bar, 10 µm. (C, D) Quantification of the number of Runx2+ cells (C) or Osterix+ cells (D) located in the bone defect beyond the proximal and distal injury sites of the centre segment at 3 dpi. N (experiments)=2, n (rays)=15. Error bars represent 95% CI. Wilcoxon matched-pairs test. (E) At 5 dpi, mineralised bone as detected by alizarin red staining has formed beyond the distal, but not the proximal injury site of the centre segment. Dashed line indicates centre segment. n=16/20 rays with distal bone formation. Scale bar, 100 µm. (F) Distribution of all positions along the proximodistal axis of centre segments, where proliferating EdU+ Zns5+ osteoblasts were observed at 2 and 3 dpi. Yellow dashed lines indicate segment border. Solid red lines, median; dashed lines, quartiles. N (experiments)=1, 2 dpi: n (rays)=13, n (cells)=306; 3 dpi: n (rays)=17, n (cells)=305; random set: n (groups)=12, n (points)=360. Kolmogorov-Smirnov test. (G) RNAscope in situ detection of bglap expression in the centre segment and the two adjacent segments distal to the bone defect at 1 dpi. Dashed lines indicate the proximal, central, and distal regions of the centre segment used for quantification. Bracket indicates dedifferentiation zone in the distal segments. Scale bar, 100 µm. (H, I) Single cell analysis of RNAscope intensity of bglap (H) or cyp26b1 (I) relative to the brightest signal in proximal, central, and distal regions of the centre segment at 1 dpi. N (experiments)=2, n (segments)=8 (bglap), 7 (cyp26b1). Error bars represent 95% CI. Holm-Sidak’s multiple comparison test. (J) Migration of bglap:GFP+ osteoblasts towards both injury sites of the centre segment. Yellow dashed lines, segment borders. Pink arrowheads indicate relocation of GFP+ osteoblasts. Asterisk indicates GFP+ osteoblasts that have entered the distal bone defect. Distal to the right. N (experiments)=4, n (segments)=95. Error bars represent 95% CI. Wilcoxon matched-pairs test. Scale bar, 100 µm. (K, L) Sequential hemiray injuries reveal no preference for osteoblast migration in distal vs proximal directions. Relocation of bglap:GFP+ osteoblasts in the centre segment (segment B) is plotted. Error bars represent 95% CI. Red arrowheads indicate time points at which the adjacent hemirays (A=proximal, C=distal) were removed. Negative relocation indicates increased distance between GFP+ cells and the joint. (K) Removal of the proximal adjacent segment A at 0 dpi, followed by removal of the distal adjacent segment C at 1 dpi. (L) Removal of the distal adjacent segment C at 0 dpi, followed by removal of the proximal adjacent segment A at 1 dpi. (M) Migration of osteoblasts into the bone defect at 3 dpi. Treatment with FK506 induces recruitment of GFP+ cells beyond the proximal injury site and increases the number of centre segments that show recruitment of GFP+ cells beyond the distal injury site. N (experiments)=1, n (segments)=56 (DMSO), 17 (FK506). Fisher’s test.

Video 2
Revascularisation of the centre segment.

Live imaging of blood flow at 2 days post injury (dpi). Yellow arrowheads indicate segment borders of the centre segment. Lower ray is not injured. Distal to the right. Scale bar, 100 µm.

In response to regular fin amputations, the initiation of osteoblast dedifferentiation, migration, and proliferation occur prior to the accumulation of a preosteoblast population in the blastema. Thus, we asked whether any of these osteoblast injury responses occur at proximal-facing injuries in the hemiray injury model, which fail to form a blastema containing preosteoblasts. At 2 and 3 dpi, proliferation of osteoblasts was observed throughout the centre segment (Figure 6F). While at 2 dpi, the median of the distribution of proliferating osteoblasts along the proximodistal axis was slightly shifted distally compared to a random distribution set, at 3 dpi proliferation was equally distributed along the segment (Figure 6F). Thus, the polarised regenerative response is not reflected by equally polarised osteoblast proliferation within the centre segment.

We next wondered whether distally-oriented bone regeneration is due to varying degrees of osteoblast dedifferentiation along the segment. We analysed the expression of bglap with single cell resolution in a proximal, central, and distal region of the centre segment at 1 dpi. Within such a distance (~300 µm), a gradual change in bglap intensity can be observed after fin amputation (Figure 1B) and in the segments distally of the distal bone defect in the hemiray injury model (bracket in Figure 6G, right panel). Yet, we could not detect differences in the extent of bglap expression between the regions of the centre segment (Figure 6G, left panel, Figure 6H), indicating that the proximal and distal injury sites induced osteoblast dedifferentiation. Similarly, cyp26b1 was upregulated along the entire segment, with no enrichment at the distal injury (Figure 6I, Figure 6—figure supplement 1B). Together, these data show that osteoblast dedifferentiation occurs to a similar extent in the bone facing the proximal and distal injury.

Intriguingly, osteoblasts in the centre segment also migrated towards both amputation planes, ultimately spreading across the entire segment (arrowheads in Figure 6J). The magnitude of migration was similar towards both wound sites (Figure 6J). This suggests that at this early time point, there is no hierarchy between the injury sites, and that osteoblasts react to an initial wound signal, which is independent of the proximodistal position of the injury. Furthermore, as osteoblasts also migrate towards the proximal site where no blastema will form, it appears that osteoblasts migrate independently of whether this will be followed by bone regeneration. To further test this, we temporally separated the creation of the two bone defects flanking the centre segment (Figure 6K and L). We first extracted only one hemiray (segment A) and analysed osteoblasts of the adjacent distal segment (segment B). Within 1 day, the distance between osteoblasts and the distal segment border increased, while proximally it decreased, revealing a directed migration of the osteoblasts towards the proximally facing injury (Figure 6K). We then removed the hemiray of segment C (the one located distally to the centre segment B) and analysed osteoblast locations in segment B 1 day later. The distance between osteoblasts and the proximal segment border further decreased, indicating that the osteoblasts continued to migrate towards the proximal injury (Figure 6K). However, at the distal side, the previously increased distance was decreased, suggesting that osteoblasts reversed their direction and migrated towards the distal injury as well (Figure 6K). This phenomenon is independent of the order of hemiray removal: when the distal adjacent hemiray (segment C) was removed first, osteoblasts migrated distally (Figure 6L). When we then removed the proximal adjacent hemiray (segment A), the distal osteoblasts continued to migrate distally, while the proximally located osteoblast migrated towards the new injury site, even though at this site no blastema will form (Figure 6L). These data strongly suggest that all bone injuries release signals that attract osteoblasts and that osteoblasts are equally likely to migrate proximally and distally. Thus, the lack of regenerative growth and bone formation at proximally-facing injuries cannot be explained by an intrinsic polarity or bias in the migration of osteoblasts on the bone matrix along the proximodistal axis of the fin.

At 2 dpi, at the distal-facing injury site, bglap:GFP+ osteoblasts of the centre segment accumulated beyond the bone matrix in the defect, where a blastema appears to form (asterisk in Figure 6J). However, no osteoblasts could be observed in the defect at the proximal side of the centre segment at 3 dpi (Figure 6J). Thus, while osteoblasts migrate along the bone matrix towards both injury sites, they exclusively migrate off the centre segment and into the defect at the distal-facing injury. In a cavity injury model, Cao et al. have shown that inhibition of the Ca2+/calmodulin-dependent phosphatase calcineurin can induce blastema formation at proximal-facing injuries (Cao et al., 2021). To analyse if such an intervention can also trigger osteoblast migration beyond the bone matrix at the proximal injury site, we treated fish with the calcineurin inhibitor FK506. Indeed, we could observe recruitment of bglap:GFP+ cells into the defect at the proximal site in FK506 treated fish (Figure 6M). Surprisingly, however, migration beyond the distal amputation site was also enhanced (Figure 6M). Therefore, to test whether calcineurin inhibition specifically enabled migration of osteoblasts into proximally-facing bone defects or whether it generally facilitated migration in all directions, we analysed osteoblast migration after regular fin amputation. FK506 treatment enhanced osteoblast migration at 1 dpa (Figure 6—figure supplement 1C), indicating that calcineurin might generally negatively regulate osteoblast migration and suggesting that it does not specifically facilitate distally directed regenerative responses.

The lack of osteoblast accumulation at proximally-facing injuries could be due to absence of chemical or mechanical cues that allow them to migrate into the defect. Alternatively, migration could be actively inhibited or osteoblasts could be eliminated at proximal injuries. One possibility would be increased osteoblast apoptosis at proximally-facing injuries. However, we could not detect any apoptotic osteoblasts at all in the first 2 days after injury (data not shown), and also at 3 dpi the number of apoptotic osteoblasts was negligible at both the proximal and distal injury sites (Figure 6—figure supplement 1D). Another possibility would be that osteoclasts interfere with osteoblast differentiation and bone formation at the proximal injury. To analyse if more osteoclasts are recruited to the proximal injury in the hemiray removal model, we analysed the distribution of cathepsinK:YFP+ cells, a marker for osteoclasts, after hemiray removal. However, osteoclasts accumulated at both wound sites by 3 dpi (Figure 6—figure supplement 1E).

Bone is thicker at the proximal base of the fin than at its distal end (Mari-Beffa and Murciano, 2010; Pfefferli and Jaźwińska, 2015). We thus wondered whether this structural heterogeneity along the fin ray could determine the different regenerative outcomes at proximal vs distal injuries. Yet, we did not observe a measurable difference in the diameter, radius, and thickness of one hemiray segment at its proximal vs distal end (Figure 6—figure supplement 1F). This suggests that structural heterogeneities of the bone along the proximodistal axis are too small at the scale of the centre segment to explain the radically different regenerative outcome at the proximal vs distal injury in the hemiray removal model.

Discussion

Generic and regeneration-specific responses of osteoblasts to injury

In this study, we interrogate the inter-relatedness of several in vivo responses of osteoblasts to injury using the zebrafish fin model. After fin amputation, osteoblasts elongate and orient along the proximodistal axis of the fin and migrate distally towards the amputation plane (Figure 7, upper panel). In addition, osteoblasts dedifferentiate, that is, they downregulate expression of differentiation markers like bglap and entpd5 and initiate expression of the preosteoblast marker runx2a. Finally, they proliferate and migrate off the bone surface to found a population of osteogenic progenitors within the regeneration blastema that drives bone regeneration (Figure 7, upper panel). Surprisingly, we find that these injury responses appear to occur largely independently of each other. In particular, osteoblast dedifferentiation does not seem to be a prerequisite for osteoblast cell shape changes and migration. These conclusions are based on two lines of evidence: first, we identified molecular interventions that can interfere with one process without affecting the others: RA and NF-κB signalling pathways regulate only osteoblast dedifferentiation, but not cell shape changes and migration, while the complement and actomyosin systems are required for osteoblast cell shape changes and migration, but not for dedifferentiation (Figure 7, upper panel). Second, using a hemiray injury model, where a proximally- and distally-facing bone injury is introduced in the same fin ray, we find that dedifferentiation, migration, and regenerative bone growth do not occur at both injuries. Osteoblasts migrate towards both sites and dedifferentiate equally in the proximity of both injuries (Figure 7, lower panel). Stunningly, however, osteoblast migration beyond the bone into the injury defect, formation of a blastema containing a preosteoblast population, and regenerative bone growth only occur at the distally-facing injury (Figure 7, lower panel). Thus, osteoblast migration and dedifferentiation represent generic injury responses that can occur independently of a regenerative response.

Model for osteoblast responses to fin amputation (upper panel) and hemiray removal (lower panel).

Osteoblast dedifferentiation and migration represent generic injury responses that are differentially regulated and can occur independently of each other and of regenerative bone growth. Upper panel: After fin amputation, osteoblasts downregulate the expression of differentiation markers. The extent depends on their distance to the amputation site, with osteoblasts close to the amputation site displaying more pronounced dedifferentiation. In addition, osteoblasts elongate and migrate towards the amputation plane and beyond to found osteogenic cells in the blastema (pink). Thus, osteoblast dedifferentiation, migration, and bone regeneration are all distally oriented. Osteoblast dedifferentiation is negatively regulated by NF-κB and retinoic acid signalling, while actomyosin dynamics and the complement system are required for directed osteoblast migration. Lower panel: In the hemiray removal model, a proximal and a distal injury are created on both sides of a remaining centre segment. The extent of osteoblast dedifferentiation is even along the centre segment, and osteoblasts migrate towards both injury sites. Yet, only at the distally-facing site, osteoblasts migrate into the bone defect, and blastema formation and bone regeneration only occur here.

The fact that dedifferentiation and migration do not always result in bone regeneration raises questions about their function. We have previously shown that experimentally enhanced dedifferentiation results in an increased population of progenitor cells in the regenerate, which, however, had detrimental effects on bone maturation (Mishra et al., 2020), indicating that dedifferentiation must be kept within certain limits. Unfortunately, currently existing tools to block dedifferentiation are either mosaic (activation of NF- κB signalling using the Cre-lox system) or cannot be targeted to osteoblasts alone (treatment with RA). Due to these limitations in our assays, we can currently not test what consequences specific, unmitigated perturbation of osteoblast dedifferentiation has for overall fin/bone regeneration. Conversely, the interventions presented here that specifically perturb osteoblast migration are limited as they act only transiently, that is, they can severely delay, but not fully block migration. Furthermore, while interference with actomyosin dynamics reduces regenerative growth, we cannot distinguish whether this is caused by the inhibition of osteoblast migration or due to other more direct effects on cell proliferation and tissue growth. Thus, an unequivocal test of the importance of osteoblast migration for bone regeneration requires different tools. However, it has been shown that fin bone can regenerate even after genetic ablation of osteoblasts, due to activation of non-osteoblastic cells which can drive de novo osteoblast differentiation (Singh et al., 2012; Ando et al., 2017). We speculate that the robustness of bone regeneration in zebrafish is enhanced by redundant mechanisms of source cell formation, which can either derive from pre-existing osteoblasts, which dedifferentiate and migrate towards the injury, or from (reserve) stem/progenitor cells. Dedifferentiation and migration in response to injuries that eventually do not trigger bone regeneration likely add to the robustness, since they prime all injuries for regeneration. Interestingly, work in planaria and zebrafish fins has previously shown that all wounds, irrespective of whether they trigger structural regeneration or result merely in wound healing, appear to activate a generic signalling response (Wurtzel et al., 2015; Owlarn et al., 2017). The differential outcome (structural regeneration vs wound healing) is rather determined further downstream and might dependent on systems that measure how much anatomy is missing (Owlarn et al., 2017). In the present work, we show that this principle also applies to injury responses that occur within osteoblasts. It appears that generic wounding-induced signals trigger the generic responses of osteoblast dedifferentiation and migration, while the decision whether these will be followed by bone regeneration is controlled by other determinants further downstream. Currently, the molecular nature of determinants that can sense the amount of missing anatomy and determine the difference between proximally and distally facing injuries remains rather mysterious.

The complement system regulates osteoblast migration in vivo

More than 20 different factors have been found to elicit a migratory response of osteoblasts in in vitro gain-of-function assays, including the activated complement peptides C5a and C3a (Dirckx et al., 2013; Thiel et al., 2018). However, confirmation of the role of many of these factors in regulating osteoblast migration in in vivo models is still sparse, largely due to the difficulty in assaying osteoblast migration independently of other phenotypes, for example, osteoblast proliferation or differentiation, in rodents in vivo. Using live zebrafish and time lapse imaging, we show that the complement system regulates osteoblast migration in vivo. Thus, the bone fracture repair defects observed in C5-deficient mice (Ehrnthaller et al., 2013) might be due to impaired recruitment of osteoblasts to the injury. The c3ar1 and c5aR1 receptors are expressed in mature fin osteoblasts, making it likely that the complement cascade directly regulates osteoblast migration. Interestingly, our data indicate that in zebrafish, both C3a and C5a act as guidance cues for osteoblasts, ensuring the recruitment of osteoblasts to the injury plane.

There is growing evidence that complement factors can modulate highly diverse processes such as cell growth, differentiation, and regeneration in various tissues. Both C3 and C5 were shown to play a role during liver regeneration in mammals, as C5−/−, C3−/− as well as C3R−/− deficient mice show impaired liver regeneration (Mastellos et al., 2001; Markiewski et al., 2004). Expression of c5aR1 is upregulated in regenerating hearts of zebrafish, axolotls, and neonatal mice, and receptor inhibition impairs cardiomyocyte proliferation (Natarajan et al., 2018). Noteworthy, in our study on zebrafish fin regeneration, inhibition of C5aR1 had no effect on osteoblast proliferation and regenerative growth. The distinct outcome of complement activation might be dependent on the location of complement factors. Intracellular C3 and C5 storage has been proposed in T cells, and the respective cleavage products C3a and C5a bind to intracellular receptors (Liszewski et al., 2013; Arbore et al., 2016). In human CD4+ T cells, intracellular C3a regulates cell survival, while activation of membrane-bound C3aR modulates induction of the immune response (Liszewski et al., 2013). In addition to the receptors, both C3 and C5 are expressed in mammalian osteoblasts (Ignatius et al., 2011b). While we lack the tools to determine the spatiotemporal distribution of the C3 and C5 precursor proteins and their activated cleavage products C3a and C5a, our expression data indicates that the majority of the precursors are produced in the liver and are distributed via the circulation. Interestingly, however, local, injury-induced transcription of c5 and the c3 paralog c3a.5 might contribute to activation of the cascade in response to fin injury as well. Intriguingly, during newt and urodele limb regeneration, C3 is expressed in blastema cells, which implies a role in mediating regeneration (Del Rio-Tsonis et al., 1998; Kimura et al., 2003). However, as during zebrafish fin regeneration osteoblasts migrate before blastema formation, a different cellular source likely produces C3 and C5. Yet, we cannot exclude that complement peptides emanating from the blastema are responsible for the migration of osteoblasts beyond the bone context at later stages, which we only observed at sites where a blastema forms. Prior to blastema formation, the injury site is covered by a wound epidermis. During newt limb regeneration, C5 was shown to be absent from the blastema but highly expressed in the wound epidermis (Kimura et al., 2003). Thus, complement activation in epidermal cells might regulate the early migration of osteoblasts to the injury.

Determinants of regeneration-specific osteoblast injury responses

Removing part of the bone within a ray results in polarised bone regeneration that is exclusively directed towards the distal end of the fin. In a recently published zebrafish fin cavity injury model, blastema markers were shown to be solely upregulated at the distal facing injury (Cao et al., 2021). In our hemiray-removal model, we analyse the proximal and distal injury of a centre segment. Thus, both injury sites are equally severed from innervation and blood supply, giving us confidence that the differential regenerative response at proximal and distal injuries is not due to differences in circulation or innervation. The distally oriented polarised regenerative response implies intrinsic mechanisms regulating diverse outcome from seemingly equal injuries (the removal of the adjacent hemiray). Our findings show that such regulators of distally- oriented regeneration seem to act downstream to or independently of regulators of other injury responses like osteoblast migration, dedifferentiation, and proliferation, that occur at all injuries. One previously suggested regulator of distally- oriented regeneration is the Ser/Thr phosphatase calcineurin (Cao et al., 2021). During zebrafish fin regeneration, inhibition of calcineurin increases regenerative growth (Kujawski et al., 2014). Importantly, the polarised, distally-oriented blastema formation in the cavity injury model can be overridden by calcineurin inhibition, which induces blastema formation also at the proximal injury (Cao et al., 2021). We also found that calcineurin inhibition can trigger the recruitment of osteoblasts into the bone defect at the proximal injury site in our hemiray removal model. Yet, we also observed increased distally-oriented osteoblast migration upon FK506 treatment, suggesting that calcineurin does not specifically regulate the polarity of injury responses along the proximodistal axis, but generally dampens responses at all injuries, including osteoblast migration and cell proliferation.

In conclusion, our findings support a model in which zebrafish fin bone regeneration involves both generic and regeneration-specific injury responses of osteoblasts. Morphology changes and directed migration towards the injury site as well as dedifferentiation represent generic responses that occur at all injuries even if they are not followed by regenerative bone formation. While migration and dedifferentiation can be uncoupled and are (at least partially) independently regulated, they appear to be triggered by signals that emanate from all bone injuries. In contrast, migration off the bone matrix into the bone defect, formation of a population of (pre-)osteoblasts, and regenerative bone formation represent regeneration-specific responses that require additional signals that are only present at distal-facing injuries. The identification of molecular determinants of the generic vs regenerative responses will be an interesting avenue for future research.

Materials and methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Genetic reagent (Danio rerio)bglap:GFP; Ola.Bglap.1:EGFPhu4008Knopf et al., 2011ZDB-ALT-110713–1
Genetic reagent (Danio rerio)osx:CreER; OlSp7:CreERT2-p2a-mCherrytud8Knopf et al., 2011ZDB-
TGCONSTRCT-100928–1
Genetic reagent (Danio rerio)hs:R to G; hsp70l:loxP DsRed2 loxP nlsEGFPtud9Knopf et al., 2011ZDB-
TGCONSTRCT-100928–2
Genetic reagent (Danio rerio)hs:Luc to IKKca BFP; hsp70l:loxP Luc-myc Stop loxP IKKca-t2a-nls-mTagBFP2-V5, cryaa:AmCyanulm12TgMishra et al., 2020ZDB-ALT-181018–2
Genetic reagent (Danio rerio)entpd5:kaede; TgBAC(entpd5a:Kaede)Huitema et al., 2012ZDB-
TGCONSTRCT-150223–1
Genetic reagent (Danio rerio)cathepsinK:YFPApschner et al., 2014
Genetic reagent (Danio rerio)hs:Luc to nYPet IĸBSR; hsp70l:loxP Luc2-myc Stop loxP nYPet-p2a-IĸBSR, cryaa:AmCyanulm15TgThis paperCre responder line to manipulate NF-ĸB signalling
AntibodyAnti-Zns5 (mouse monoclonal)Zebrafish resource center, Eugene USARRID:AB_10013796IF (1:300)
AntibodyAnti-Runx (mouse monoclonal)Santa CruzRRID:AB_1128251IF (1:300)
AntibodyAnti-Osterix (rabbit polyclonal)Santa CruzRRID:AB_831618IF (1:300)
Commercial assay or kitRNAscope Multiplex Fluorescent kit v2ACD Bio-techneCat# 323,100
Commercial assay or kitApopTag Red In Situ Apoptosis Detection KitMerckCat# S7165
Commercial assay or kitEdU-Click 647 kitBaseclick GmbHCat# BCK-EdU647
Commercial assay or kitLunaScript RT SuperMIX KitNEBCat# E3010
Chemical compound, drugCytochalasin DCalbiochemCat# 250,255
Chemical compound, drug(-)-blebbistatinSigmaCat# B0560
Chemical compound, drugW54011EnzoCat# ENZ-CHM122-0001
Chemical compound, drugSB290157CaymanCat# 15,783
Chemical compound, drugPMX205TocrisCat# 5196
Chemical compound, drugRetinoic acidSigmaCat# R2625
Chemical compound, drugNocodazoleSigmaCat#M1404
Chemical compound, drugFK506MerckCat# F4679
Chemical compound, drug(Z)–4-HydroxytamoxifenSigma AldrichCat# H7904
Software, algorithmFijiSchindelin et al., 2012
Software, algorithmGraphPad Prism 9GraphPad software

Animals

All procedures involving animals adhered to EU directive 2010/63/EU on the protection of animals used for scientific purposes and were approved by the state of Baden-Württemberg (Project numbers 1193 and 1494) and by local animal experiment committees. Fish of both sexes were used. Housing and husbandry followed the recommendations of the Federation of European Laboratory Animal Science Associations (FELASA) and the European Society for Fish Models in Biology and Medicine (EUFishBioMed) (Aleström et al., 2020).

The following pre-existing transgenic lines were used: bglap:GFP (Ola.Bglap.1:EGFPhu4008; Knopf et al., 2011), osx:CreER (Ola.Sp7:CreERT2-p2a-mCherrytud8; Knopf et al., 2011), entpd5:kaede (TgBAC[entpd5a:Kaede]; Huitema et al., 2012), cathepsinK:YFP (Apschner et al., 2014), hs:R to G (hsp70l:loxP DsRed2 loxP nlsEGFPtud9; Knopf et al., 2011), hs:Luc to IKKca BFP (hsp70l:loxP Luc-myc Stop loxP IKKca-t2a-nls-mTagBFP2-V5, cryaa:AmCyanulm12Tg; Mishra et al., 2020). To create the line hsp70l:loxP Luc2-myc Stop loxP nYPet-p2a-IκBSR, cryaa:AmCyanulm15Tg, in short hs:Luc to nYPet IκBSR, the following elements were assembled by Gibson assembly and restriction-based cloning methods: MiniTol2 inverted repeat, attP site, zebrafish hsp70l promoter, loxP, firefly luciferase 2, 6× myc tag, ocean pout antifreeze protein polyA signal, loxP, nYPet, p2a, IκBSR (IkappaB super-repressor), SV40 polyA signal, zebrafish cryaa promoter, AmCyan, SV40 polyA signal, MiniTol2 inverted repeat. IkBSR is a mutated version of mouse IKappa B alpha (Entrez gene, Nfkbia A1462015), described in Van Antwerp et al., 1996, whose inhibitory N-terminal Serine residues 32 and 36 are mutated to Alanine. Phosphorylation sites in the C-terminal PEST domain are also mutated: Serines 283, 288, and 293 are converted to Alanine; Threonines 291, 296 also to Alanine, and Tyrosine 302 to Aspartic acid. Tol2-mediated transgene insertion was used to create a stable transgenic line. One subline was selected based on widespread expression after heat shock in the adult fin and efficient recombination in embryos when crossed with a ubiquitin promoter-driven Cre driver line (unpublished).

To achieve optimal expression levels of the osx:CreER transgene, we activated its expression in adult fins as described previously (Mishra et al., 2020). Briefly, fish were amputated and allowed to regenerate for 8 days, at which timepoint mineralised bone has formed in the regenerate. We then reamputated through the mineralised part of the regenerate and analysed the stump of this second amputation.

Fin amputations and hemiray removal

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Adult zebrafish were anaesthetised with 625 µM tricaine, and the caudal fins were amputated through the second segment proximal to the bifurcation. For hemiray removal, small surgical blades were used to cut through four joints proximal to the bifurcation in the second ray from ventral and dorsal, and fine tweezers were used to remove the upper, left hemiray (facing towards the experimenter) of the segments located proximal and distal of one central segment. For analysis of migration and dedifferentiation, one hemiray segment was removed on either side. For analysis of proliferation and Runx2 and Osterix expression, two hemiray segments were removed on both sides. For alizarin red staining at 5 dpi, three hemiray segments were removed. Fish were allowed to regenerate at 27–28.5°C.

qRT-PCR

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Total RNA was extracted from fins using RNeasy Mini Kit (Qiagen, 74104). For liver RNA, six adult livers were pooled for each replica, for non-injured fins, four whole fins were pooled, and for 6 hpa fin samples, segment –1 and segment 0 from 15 fins were pooled. cDNA synthesis was carried out using LunaScript RT SuperMIX Kit (New England Biolabs, E3010L) with random primers. Quantitative real time PCR was performed using Luna Universal qPCR Master Mix (NEB, M 3003 L) with the following primers:c5 (ZDB-GENE-120510–2, fw: CCGGTCACTACAGTCAATGGT, rv: CGTTGAGGGAGTAAACACGC), c5ar (ZDB_GENE-190430–1, fw: TCGGATGCCAAACTCAGTGA, rv: CATGAGGATGGGAAGGGACA), c3a.1 (ZDB-GENE-990415–35, fw: GGAGATGGACCAGAGTGTGT, rv: ATCAATCTCCTCCCACAGCC), c3a.2 (ZDB-GENE-990415–36, fw: CGGTACACAAACACCCCTCT, rv: GTCTTCCTCGTCGTTCTCTTGTT), c3a.3 (ZDB-GENE-990415–37, fw: TGTTGATGAATCTAAGCGCTTGA, rv: CGGAGTCTTGTTTCAGGTCC), c3a.4 (ZDB-GENE-140822–3, fw: TGGTGGCTGTGGATAAAGGT, rv: CTGTGCAGCCAGTGTCATG), c3a.5 (ZDB-GENE-090311–30, fw: ACCAAGAATCTCTGACTGTGGA, rv: CCGCCATGCTGATCTTCTTC), c3a.6 (ZDB-GENE-041212–2, fw: TCTGGAAGGTGGTCACAAGA, rv: TCGATGCTAACCGTCAGACT), and c3aR1 (ENSDARG00000031749, fw: CATGCTGGCTGTTATCGTGG, rv: AACACATACAGGACGGGGTT). Relative quantification was performed using the ΔΔCt method, and the Ct values of each gene were normalised to the arithmetic mean of two housekeeping genes, actb2 (ZDB-GENE-000329–3, fw: ACGATGGATGGGAAGACA, rv: AAATTGCCGCACTGGTT) and hatn10 (ZDB-NUCMO-180807–7, fw: TGAAGACAGCAGAAGTCAATG, rv: CAGTAAACATGTCAGGCTAAATAA). Each biological sample was run in technical duplicates and results were averaged. qPCRs and statistics were performed on two biological replicates for liver samples, three biological replicates for non-injured fin samples, three biological replicates for injured fin samples, and four biological replicates for cells isolated by FACS (Fluorescence Activated Cell Sorting).

Fluorescence Activated Cell Sorting of osteoblasts

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Differentiated osteoblasts were isolated at 1 dpa from segment 0 and segment –1 of adult bglap:GFP transgenics using FACS as described by Lee et al., 2020. A total of 15 fins were pooled for each of the 4 replicates. In brief, samples were treated with 1× TrypLE Express enzyme (Thermo Fisher, 12604013) at 37°C for 30 min with gentle agitation, washed with cold Dulbecco’s phosphate buffered saline (DPBS) buffer (Thermo Fisher, 14190136), and further dissociated into a single cell suspension with 0.25 mg/mL Liberase DL (Merck, 5401160001) in enzyme-free Cell Dissociation Buffer (Thermo Fisher, 13151014) at 37°C for 30 min with gentle agitation. The dissociated single cell suspension was washed with cold DPBS buffer and pelleted by centrifugation at 500× g for 3 min at 4°C. Cells were resuspended in cold DPBS buffer with 2% fetal bovine serum and filtered through a 70 μm sample preparation filter (pluriSelect, 43-10070-46) to remove cell aggregates. The FACS Aria II flow cytometer was used to separate GFP+ and GFP− populations from single-cell suspensions. In order to define GFP background levels, wildtype non-transgenic fish were used. Forward and side scatter parameters were applied to exclude debris and doublets. Dead cells were excluded by staining with the cell-impermeable DNA-dye 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI, Sigma, 32670) and sorting for DAPI-negative cells. To further discriminate between GFP+ anuclear cell debris and intact cells, we stained with the cell-permeable DNA dye DRAQ5 (Cell Signalling, 4084 S) and sorted for DRAQ5+ cells. Therefore, DAPI−/DRAQ5+/GFP+ cells were sorted, and the same number of DAPI−/DRAQ5+/GFP− cells from each sample were also sorted using a random GFP gate to serve as control group.

Pharmacological interventions

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10 µl of the following drugs (in PBS) were injected IP using a 100 µl Nanofil syringe (World Precision Instruments NANOFIL-100): blebbistatin 750 nM, cytochalasin D 30 µM, W54011 10 µM, SB290157 10 µM, PMX205 10 µM. Fish were injected once daily, starting 1 hr prior to injury. For the following treatments, fish were immersed in fish system water containing the following drugs: RA 5 µM, nocodazole 5 µM, FK506 3 µM. Fish were set out in drug solution 1 hr prior to injury, and solutions were changed daily. For the entire duration of the experiment, fish were kept in an incubator at 25°C in the dark at 1 fish/100 ml density in fish system water. Negative control groups were injected or soaked, respectively, with the corresponding DMSO concentration in PBS or fish system water.

Whole mount immunohistochemistry

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Fins were fixed overnight with 4% PFA at 4°C. After 2 × 5 min washes with PBTx (1× PBS with 0.5% TritonX 100) at RT, fins were transferred to 100% acetone, rinsed once, and kept for 3 hr at −20°C. Fins were transferred to PBTx, washed 2 × 5 min, 2 × 15 min, and incubated for 30 min at RT, followed by blocking in 1% BSA in PBTx for at least 1 hr at RT. Primary antibodies were diluted to 1:300 in blocking solution and fins were incubated overnight at 4°C. The next day, fins were washed several times with PBTx and incubated with secondary antibodies at 1:300 dilution in PBTx overnight at 4°C. Fins were mounted in Vectashield (Vectorlabs, H-1000). For anti-Runx2 staining, fins were fixed with 80% methanol/20% DMSO overnight at 4°C and rehydrated by a graded series of methanol in PBTx (75, 50, 25%) for 5 min each. Fins were washed in PBTx for 30 min, after which fins were processed following the above-mentioned protocol. Primary antibodies used in this study were: mouse anti-Zns5 (Zebrafish International Resource Center, Eugene, OR, USA, RRID:AB_10013796), mouse anti-Runx (Santa Cruz sc-101145, RRID:AB_1128251), and rabbit anti-Osterix (Santa Cruz sc-22536-R, RRID:AB_831618).

RNAscope whole mount in situ hybridisation

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For detection of RNA transcripts in whole mount fins, the RNAscope Multiplex Fluorescent Reagent Kit v2 (ACD Bio-techne, 323100) was used with the following probes: bglap-C1 (ACD Bio-techne, 519671), cyp26b1 (ACD Bio-techne, 571281-C2), entpd5 (ACD Bio-techne, 820491-C4), runx2a (ACD Bio-techne, 409521-C2), and c5aR1 (ACD Bio-techne, 859561-C2). Fins were fixed overnight in 4% PFA at 4°C. The next day, fins were washed with PBT (1× PBS with 0.1% Tween) 3 × 5 min each. After incubation in RNAscope hydrogen peroxide for 10 min, fins were rinsed 3× with PBT, treated with RNAscope protease plus for 20 min, and rinsed 3× with PBT. All these steps were performed at RT. Probes and probe diluent were prewarmed to 40°C and cooled down to RT before use. The fins were incubated with probes overnight at 40°C. The next day, fins were washed 3× with 0.2× SSCT, 12 min each at RT. All the steps mentioned from here on were followed by such washing steps. Fins were post fixed with 4% PFA for 10 min at RT and incubated with AMP1, AMP2, and AMP3 for 30, 15, and 30 min, respectively, at 40°C. To develop the signal, fins were incubated with the corresponding RNAscope Multiplex FL v2 HRP for 15 min at 40°C. TSA-fluorophore (Perkin Elmer NEL701A001KT) was used in a 1:1500 dilution in TSA buffer (RNAscope kit). Fins were incubated for 15 min at 40°C.

Proliferation assay

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To analyse cell proliferation, 5-ethynyl-2′-deoxyuridine (EdU)-Click 647 kit (baseclick GmbH BCK-EdU647) was used. Fish were IP injected with 10 µl of 10 mM EdU in PBS using a 100 µl Nanofil syringe (World Precision Instruments NANOFIL-100). In amputation assays, fish were injected as indicated in the figures. In the hemiray removal assay, fish were injected once at 1 dpi for evaluation at 2 dpi, and at 1 and 2 dpi for evaluation at 3 dpi. Fins were fixed with 4% PFA overnight and EdU labelling was performed according to the manufacturer’s protocol. If applicable, EdU labelling was followed with AB staining following the immunohistochemistry protocol, or RNAscope in situ following the hybridisation protocol.

Apoptosis assay

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To analyse apoptosis, the ApopTag Red In Situ Apoptosis Detection Kit (Merck, S7165) was used. Fins were fixed overnight in 80% Methanol/20% DMSO at 4°C. The next day, fins were rehydrated through a Methanol/PBTx (PBS +0.5% TritonX 100) series (75, 50, 25%; 5 min each) and permeabilised with acetone for 3 hr at –20°C. Fins were transferred to PBTx, washed 2 × 5 min, and incubated for 30 min at RT, followed by blocking in 1% BSA in PBTx for at least 1 hr at RT. Mouse anti-Zns5 (Zebrafish International Resource Center, Eugene, OR, USA, RRID:AB_10013796) was diluted to 1:300 in blocking solution and fins were incubated overnight at 4°C. The next day, fins were washed several times with PBTx and transferred to superfrost slides. Using a hydrophobic pen, a circle was drawn around the fins. Fins were first equilibrated with 75 µl equilibration buffer (ApopTag kit) for 10 s at RT, followed by incubation in 55 µl TdT enzyme (ApopTag kit) in a humidified chamber at 37°C for 1 hr. The reaction was stopped by several washes with Stop/Wash buffer (ApopTag kit), followed by an incubation in Stop/Wash buffer for 10 min at RT. Fins were washed 3 × 1 min with PBS and incubated in anti-DIG-Rhodamine diluted 1:2000 in blocking solution (ApopTag kit) and secondary antibody (1:300) overnight at 4°C. The next day, fins were washed 6 × 20 min at RT and mounted with Vectashield (Vectorlabs, H-1000).

Alizarin red S staining

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In vivo staining with alizarin red S (ARS; Sigma-Aldrich A3757) was performed as previously described (Bensimon-Brito et al., 2016). Briefly, fish were stained in 0.01% ARS in system water for 15 min at RT, washed 3 × 5 min in system water, immediately anaesthetised, and imaged.

Cre-lox recombination and heat shocks

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Adult double transgenic fish (OlSp7:CreERT2-p2a-mCherrytud8; hsp70l:loxP DsRed2 loxP nlsEGFPtud9, OlSp7:CreERT2-p2a-mCherrytud8; hsp70l:loxP Luc2-myc Stop loxP nYPet-p2a-IκBSR, cryaa:AmCyanulm15Tg and OlSp7:CreERT2-p2a-mCherrytud8; hsp70l:loxP Luc-myc Stop loxP IKKca-t2a-nls-mTagBFP2-V5, cryaa:AmCyanulm12Tg) were IP injected with 10 µl of 3.4 mM 4-OH tamoxifen (4-OHT) in 25% ethanol once daily for 4 days. Fish were heat shocked twice (at 24 hr and 3 hr prior to harvest) at 37°C for 1 hr after which water temperature was returned to 27°C within 15 min.

Imaging

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Images in Figures 1F, G, 3C, G, 6J, Figure 5E; Figure 5—figure supplement 4A, B and Figure 6—figure supplement 1E were acquired with a Leica M205FA stereo microscope and display live fluorescence of fluorescent proteins. High-resolution optical sections were obtained with a Leica SP8 confocal microscope or a Zeiss AxioObserver 7 equipped with an Apotome and processed using Fiji (Schindelin et al., 2012). Images in Figures 1B,C and D,2A, D and 4A, 6B,E,G; and Figure 1—figure supplement 1A, B; Figure 2—figure supplement Figure 2—figure supplements 1A and 2A,B, Figure 5—figure supplement 1A; Figure 6—figure supplement 1B,D and Video 1 are z-projections. Figure 5A and Figure 5—figure supplement 2D show single z-planes. Video 2 was recorded with a Leica M205FA stereo microscope. Movie annotations were added using the Annotate_movie plugin (Daetwyler et al., 2020).

Cell morphology quantification

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To quantify osteoblast cell shape and orientation, the transgenic line bglap:GFP in combination with Zns5 AB labelling was used. Osteoblasts of the outer layer of one hemiray (facing the objective in whole fin mounting) were imaged and analysed. As Zns5 localises to the plasma membrane of all osteoblasts, the combination of both markers provides solid definition of single cell outlines. All GFP+ Zns5+ cells with such a defined outline within an analysed segment were included into the analysis, and cells along the whole proximodistal axis of a segment were measured. In the transgenic intervention studies, mCherry is expressed under the osx promoter and was used as cytosolic labelling of osteoblasts. Using Fiji (Schindelin et al., 2012), the longest axis of a FP+ Zns5+ cell was measured as maximum length, the short axis as maximum width, and the ratio calculated. Simultaneously, the angle of the maximum length towards the proximodistal ray axis was measured for angular deviation. All measurements were performed manually, with the analyst being blinded.

Bulk migration assay

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To quantify osteoblast migration, fins of bglap:GFP transgenic fish were imaged with a GFP filter and in brightfield at 0 and 1 dpa with a Leica Stereomicroscope M205FA. Using Fiji (Schindelin et al., 2012), a threshold was set for the GFP signal to exclude background fluorescence and to select the bulk of GFP+ cells. The thresholded image was merged with the corresponding brightfield image and the distance between GFP+ cells and the joint at the ventral-distal centre of the segment was measured. For each fin, the second and third rays in the dorsal lobe of the fin were analysed. Statistical analysis was performed on the difference in distance between 0 and 1 dpa, while for illustration the change at 1 dpa is plotted, with migration across the whole distance to the joint counting as 100% migration.

RNAscope quantification

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For quantification of RNAscope signals, optical sections were acquired with a Zeiss AxioObserver 7 microscope equipped with an Apotome, using identical imaging settings within one experiment. To analyse bglap intensity along the proximodistal axis, subsequent ROIs (segment height × 50 µm width) were analysed (joints excluded), and intensity was normalised for each ray. For spatial resolution of single osteoblast intensity, segment lengths were normalised and x-location of osteoblasts grouped into proximal (0.0–0.2 normalised segment length), central (0.4–0.6), and distal (0.8–1.0) regions. For categorising bglap expression of single cells, osteoblasts were grouped into three classes based on bglap expression intensity using a look-up table (LUT). LUT was designed so that the ‘high’ threshold (pixel values 192–255) corresponds to the expression levels in segment –2 and to further robustly discriminate between ‘low’ (32-127) and ‘medium’ (128-191) cells.

Regenerative growth quantification

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Regenerative growth was analysed in brightfield images taken at 3 dpa. For each fin, the length of the second and third dorsal ray regenerate was measured from the amputation plane to the distal tip and the average calculated. For testing significance of growth differences between drug treatments and control fish, the data were box cox transformed using the formula xλ-1λ with λ=1.5, which we determined previously (Mishra et al., 2020).

Statistical analysis

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Fish were randomly allocated into experimental groups. Statistical analyses were performed using GraphPad Prism (GraphPad software, LCC). For proliferation analyses in the hemiray removal model, a random distribution set was generated with MS Excel (Microsoft Corporation). Percentage data were arcsin transformed for statistical analysis. For experiments where we did not observe significant differences between experimental groups, if applicable we used the following logic to support statements that no differences exist or that these are not biologically relevant: the smallest difference that would have been significant was calculated based on the effect size and the SDs and sample sizes of the experimental groups. Using G × Power (Faul et al., 2009) the effect size was computed with these parameters: t-test, tails=2, α=0.05, power (1−β)=0.8, and the sample size of both groups. For non-normal distributed data sets, data was log-transformed. These calculated smallest significant differences and the observed differences are reported in the figure legends. If the calculated smallest significant difference is smaller than significant differences we observed with similar assays in other experiments, we conclude that the experiment would have had enough statistical power to detect a similar effect size.

Data availability

All data generated or analysed during this study are included in the manuscript and supporting file.

References

    1. Del Rio-Tsonis K
    2. Tsonis PA
    3. Zarkadis IK
    4. Tsagas AG
    5. Lambris JD
    (1998)
    Expression of the third component of complement, C3, in regenerating limb blastema cells of urodeles
    Journal of Immunology (Baltimore, Md 161:6819–6824.
    1. Florian S
    2. Mitchison TJ
    (2016) Anti-Microtubule Drugs
    Methods in Molecular Biology (Clifton, N.J.) 1413:403–421.
    https://doi.org/10.1007/978-1-4939-3542-0_25

Decision letter

  1. Céline Colnot
    Reviewing Editor; INSERM U955, UNIV PARIS EST CRETEIL, France
  2. Didier YR Stainier
    Senior Editor; Max Planck Institute for Heart and Lung Research, Germany
  3. Céline Colnot
    Reviewer; INSERM U955, UNIV PARIS EST CRETEIL, France

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Decision letter after peer review:

Thank you for submitting your article "Zebrafish fin regeneration requires generic and regeneration–specific responses of osteoblasts to trauma" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, including Céline Colnot as the Reviewing Editor and Reviewer #1, and the evaluation has been overseen by Didier Stainier as the Senior Editor.

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

Essential revisions:

1) Additional experimental details should be provided to demonstrate the absence of biased selection of cells chosen to be incorporated into the statistical analyses. A clearer picture of how the authors obtained their final numbers starting from the original image data is needed.

2) Additional sets of marker genes (in addition to bglap:GFP downregulation) and long time–lapse imaging of the reporter (preferably with histone–GFP) would be helpful to further substantiate the claims on "dedifferentiation" and "migration" of osteoblast populations. Additional characterization of bglap:GFP+ cells would be also helpful to determine if these cells also include osteoblast precursor cells.

3) Additional results are needed to support the claim that the complement system is required for injury–induced directed osteoblast migration including expression analyses of the central complement components C3 and C5 at the amputation site and showing whether c5ar1 and c3ar are solely expressed in osteoblasts, or rather broadly within tissue lining the hemirays.

4) It is unclear whether each step of cell elongation and alignment, cell migration, cell dedifferentiation and regenerative response, is required for fin regeneration following amputation. In the absence of direct evidence for the requirement of migration or dedifferentiation for the overall success of fin regeneration, the limitations should be more clearly stated, and the title modified.

5) The word trauma should be clarified as the authors do not provide evidence for a non–healing phenotype in their trauma model.

6) The conclusions referring to the relevance of the work to bone regeneration should be toned down and more focused on fin regeneration.

7) The manuscript is overall well written but could be improved to be less technical to help the readers, and the discussion should be reduced in length.

Reviewer #1 (Recommendations for the authors):

To determine whether each step of cell elongation and alignment, cell migration, cell dedifferentiation and regenerative response, is required for fin regeneration following amputation the authors discuss p14 that other tools will be needed. However, later time points could be assessed in this study, not only to show the long–term effects of the treatment on healing but also to show how healing eventually resolves in the two models used here. Did the authors assess the long–term effects of inhibitor treatments on the regeneration process? Does blocking the complement system or actomyosin prevent regeneration? Are both generic (migration, dedifferentiation) and regenerative growth required for regeneration? Since there are limitations in the approaches used, the authors should modify the title and replace the word "requires". The use of the term "require" should be reconsidered also in the abstract and throughout the manuscript if the requirement for healing or a specific process is not demonstrated.

Can the authors provide evidence of the source of C3 and C5, the central components of the complement system during fin regeneration? Figure 4 shows RNAscope in situ detection of c5aR1. Analysis of c3 and c5 using the same technique should be included.

Regarding the trauma model, it is unclear whether the gaps on each side of the remaining segment eventually heal. Does the gap on the proximal side of the remaining segment heal via regeneration occurring on the distal end of the next segment? The term "trauma" should be replaced by another term. Further, the term "trauma" appears to be used in the first paragraph of the discussion when referring to amputation and in other parts of the discussion when referring to bone injury.

The Discussion section is very long and there are many redundancies with the Results section. The main findings and conclusions could be stated more briefly, and the discussion points presented in a more concise way. As indicated in the public review, the last paragraph of the discussion should be focused on "fin regeneration" and avoid concluding on "bone regeneration" and "bone formation".

Reviewer #2 (Recommendations for the authors):

– Page 4, line 6: I am not sure if "exoskeletal" is by definition the right term in this context.

– What is the p–value in Figure 2A (left) between the width/length ratio of cytochalasin–treated osteoblasts from segments –1 and –2? It looks as if the cells of segment –1 are somewhat elongated even though they have been exposed to the drug. Could the authors please comment on this?

– The red labeling of data points for the drug–treated conditions in Figure 2A, B and C might be difficult to be detected by color–blind readers. This should be adjusted accordingly.

– An image showing cyp26b1 RNAscope in situ hybridization on which the graphs of Figure 5H are based should be shown somewhere – maybe in the dedicated Excel sheet of raw data.

– Suggestion for a further experiment (not required for acceptance from my side!!!): I think it would be nice to learn whether C3a and C5a are sufficient to drive directed osteoblast migration along bony elements. To test that one could try to implant a bead loaded with recombinant C3a or C5a somewhere along a fin ray to see whether this ectopic signaling source attracts osteoblasts. Would this be doable? Recombinant complement proteins appear to be commercially available.

Reviewer #3 (Recommendations for the authors):

1. Title:

– I feel that the current title does not capture the essence of the authors' major findings.

2. Abstract:

– The abstract is oversimplified and overstated, omitting important details. The authors should clearly state that they used a bglap:GFP strain to support their conclusions on osteoblast differentiation and migration.

3. Introduction:

– The authors make significant statements on osteoblast dedifferentiation. This is based on a naïve assumption that bglap:GFP is solely active in mature osteoblasts. However, it is clear from the authors' data that a substantial fraction of GFP+ cells are in proliferation (Figure 4 S1B). It seems that bglap:GFP is also expressed by osteoblast precursors. Specificity of bglap:GFP to mature osteoblasts is a major concern.

– Page 5, Line 5: Osteoblasts do not migrate to the resorbed sites. Osteoblast precursors (or pre–osteoblasts) do. Thiel 2018 review article does not appear to be particularly supportive.

4. Results:

– Page 6, Line 14: Perhaps the authors should explain in more detail about the kaedeGreen to Red system, and why they needed this approach.

– Page 7, Line 13–16: I do not understand why the authors interpret these data as directed active migration behavior towards the injury. What they showed here is merely cell shape changes.

– Page 7, Line 19: This data could be alternatively interpreted as the upregulation of bglap:GFP by invading cells (not previously bglap+), rather than the invasion of bglap:GFP+ cells. It would be essential to determine the half–life of GFP to assume cell invasion. The authors would need a more long–lasting histone–bound GFP reporter strain to conclusively define cell invasion.

– Page 8, Line 3–16: This paragraph is very difficult to read because of long sentences. They should simplify and rephrase the sentences.

– Page 9, Line 1: The authors should move the bglap RNAscope in situ data to the main figure, as this is an important piece of information.

– Page 9, Line 6: I don't understand why reduced bglap mRNA can be interpreted as osteoblast dedifferentiation.

– Overall, the authors would need live–cell imaging and tracking to make these conclusions.

– Page 10, Line 7: If the authors think that a bglap:GFP strain can be used as live reporters for osteoblast dedifferentiation, that would significantly compromise the scientific rigor of the study, as it would not support any conclusions regarding cell migration and invasion.

– Page 10, Line 20: Osteoblast migration should not be mixed up with cell shape changes.

– Page 11, Line 12: The authors did not show data for osteoblast differentiation/dedifferentiation in their genetic intervention studies.

– Page 11, Line 19: An alternative explanation is that retinoic acid and NF–kB signaling simply promote osteoblast differentiation, without anything to do with dedifferentiation.

– Page 12, Line 1–2: These conclusions are stretched and not supported by experimental data. The argument for the decoupling of dedifferentiation and migration is not well developed.

– Page 13, Line 8: As many as 20% of bglap:GFP+ cells are EdU+ therefore in proliferation. How would the authors explain this?

5. Discussion:

– The authors should clearly indicate their definition of osteoblasts in this study (i.e. bglap:GFP+ cells).

– Osteoblast dedifferentiation should be rephrased as reduced bglap expression.

– Page 18, Line 19: Osteoblast differentiation, instead of osteoblast dedifferentiation.

https://doi.org/10.7554/eLife.77614.sa1

Author response

Essential revisions:

1) Additional experimental details should be provided to demonstrate the absence of biased selection of cells chosen to be incorporated into the statistical analyses. A clearer picture of how the authors obtained their final numbers starting from the original image data is needed.

Osteoblasts line the bony hemirays on the inner and outer surface (see Figure 1A), and for quantifications of osteoblast morphology, we analysed the osteoblasts of the outer layer of one hemiray (the hemiray facing the objective in whole mount imaging). For measurement of morphology, we used a transgenic line expressing a fluorescent protein (FP) in osteoblasts in combination with Zns5 antibody labelling. Zns5 is a pan-osteoblastic marker which localizes to the cell membrane. Therefore, combination of a cytosolic FP labelling with the membrane labelling by Zns5 provides solid definition of single cell outlines. For general morphology studies and drug intervention studies, we used bglap:GFP transgenics. In the transgenic intervention studies (manipulation of NF-κB signalling), mCherry is expressed together with CreERT2 under the osterix promoter and used as cytosolic labelling of osteoblasts. Our analyses are always based on segments, e.g. we present data for segments 0, -1, 2. Within these segments all FP+ Zns5+ cells were included into the analysis, and cells along the whole proximodistal axis of a segment were measured. Measurements were performed manually, and the analysist was blinded. With these set-ups, not only a fraction but all FP+ Zns5+ osteoblasts present in those segments that we analysed were included into the analysis, and thus no selection was necessary that could have introduced bias. As suggested by Reviewer #2, we have added representative sample images to the accompanying Excel sheets of raw data for the dedicated experiments. Within these, the axes along which the measurements have been performed are indicated. We have expanded the description of the analysis in the method section. It now reads on page 36 as follows:

“To quantify osteoblast cell shape and orientation, the transgenic line bglap:GFP in combination with Zns5 AB labelling was used. Osteoblasts of the outer layer of one hemiray (facing the objective in whole fin mounting) were imaged and analysed. As Zns5 localizes to the plasma membrane of all osteoblasts, the combination of both markers provides solid definition of single cell outlines. All GFP+ Zns5+ cells with such a defined outline within an analysed segment were included into the analysis, and cells along the whole proximodistal axis of a segment were measured. In the transgenic intervention studies, mCherry is expressed under the osx promoter and was used as cytosolic labelling of osteoblasts. Using Fiji (Schindelin et al., 2012), the longest axis of a FP+ Zns5+ cell was measured as maximum length, the short axis as maximum width, and the ratio calculated. Simultaneously, the angle of the maximum length towards the proximodistal ray axis was measured for angular deviation. All measurements were performed manually, with the analyst being blinded.”

2) Additional sets of marker genes (in addition to bglap:GFP downregulation) and long time–lapse imaging of the reporter (preferably with histone–GFP) would be helpful to further substantiate the claims on "dedifferentiation" and "migration" of osteoblast populations. Additional characterization of bglap:GFP+ cells would be also helpful to determine if these cells also include osteoblast precursor cells.

To address these comments, we have performed several additional experiments as described below. In addition, we would like to refer the reviewers to our previous papers, where we have analysed the process of osteoblast dedifferentiation (Knopf et al., Dev Cell 2011, doi: 10.1016/j.devcel.2011.04.014; Geurtzen et al., Development 2014, doi: 10.1242/dev.105817; Mishra et al. Dev Cell 2020, doi: 10.1016/j.devcel.2019.11.016). Using transgenic reporters and immunofluorescence we have shown in these previous papers that osteoblasts in the non-injured fin express the differentiation marker Bglap but not the pre-osteoblast marker Runx2 (and are thus by our definition differentiated), and that these cells do not only downregulate Bglap but also upregulate Runx2 in the process of dedifferentiation.

However, we have refined and expanded our methods and are now able to determine the expression patterns of markers of the osteoblast differentiation status with single cell resolution using RNAScope in situ hybridization. We think that these data are even more conclusive than those we previously published, since they don’t rely on transgenic reporters, which might not fully reflect the endogenous expression, and are not affected by issues of antibody specificity. The results that we have now obtained with this method fully support our previous conclusions.

Already in the previous version of the manuscript we have shown that endogenous bglap is strongly expressed in segment -2, (the segment that does not respond to fin amputation and thus represents the non-injured state), while it is downregulated in a graded manner in segment -1 and segment 0 (the segments where dedifferentiation happens). These data were possibly somewhat hidden in the Supplement, we have now moved them to the re-designed Figure 1B. In addition to bglap, we can now show that entpd5, a gene required for bone mineralization, is strongly expressed in osteoblasts of segment -2, while it is massively downregulated in segment -1 and segment 0. Thus, entpd5 is another differentiation marker whose loss characterizes osteoblast dedifferentiation. These new data can be found in Figure 1C. Importantly, we can confirm by RNAScope that the pre-osteoblast marker runx2a is absent in mature segments but is upregulated in segment 0 and segment -1 at 1 dpa (new data in Figure 1 —figure supplement 1A). Similarly, cyp26b1, an enzyme shown to regulate dedifferentiation, is upregulated in segment 0 and segment -1, but is not expressed in segment -2 (new data in Figure 1 —figure supplement 1B). Furthermore, we have repeated all experiments where we have previously quantified dedifferentiation upon experimental interventions using downregulation of bglap:GFP (actomyosin inhibition, retinoic acid treatment, complement inhibition). We now can fully confirm the previous conclusions using the more rigorous quantification of dedifferentiation using RNAScope analysis of endogenous bglap levels. We have replaced all bglap:GFP data with the new bglap RNAScope data. These new data are found in Figure 3F, Figure 3 —figure supplement 1A, Figure 4B and Figure 5F.

To further support our conclusion that osteoblasts do not only dedifferentiate, but also migrate, we performed time-lapse imaging using a transgenic line expressing the photoconvertible protein kaede in osteoblasts (entpd5:kaede). Local photoconversion of only the proximal half of a segment allowed us to trace these photoconverted osteoblasts. This revealed that converted cells appear in the distal part of the segment within 1 dpa, which can only be explained by relocation of the cells. These new data can be found in Figure 1F.

Our main tool to monitor osteoblast migration is the bglap:GFP transgenic line. Although bglap expression is downregulated during osteoblast dedifferentiation and thus also GFP levels eventually drop in the transgenic line, we can nevertheless use this line to trace osteoblasts, since GFP protein persists for up to three days in cells that shut down endogenous bglap and also bglap:GFP transgene transcription. While we have already shown this previously (Knopf et al., Dev Cell 2011, doi: 10.1016/j.devcel.2011.04.014), we have now also used RNAScope to confirm this. We analysed the expression of GFP on protein and RNA level in the bglap:GFP line. In bglap:GFP fish, in a mature segment in non-injured fins the regions close to the joints are devoid of cells expressing GFP (Figure 1G). Yet after amputation, we observe GFP+ cells in this distal part of segment -1 (Figure 1G, D). RNAscope in situ shows that these GFP+ cells are negative for gfp RNA (new data in Figure 1D). Thus, the observed fluorescence is due to the persistence of the GFP protein and not due to a potential upregulation of the transgene, which confirms that these GFP+ cells have migrated from the centre of the segment.

To confirm the specificity of the bglap:GFP line for mature osteoblasts, we performed immunofluorescence against Runx2 on 7 dpa regenerates, at a stage where blastema proliferation at the distal tip of the regenerate produces new osteoblast progenitors, while in more proximal (older) regions osteoblasts have already started to differentiate and new bone matrix has formed. We found that Runx2 is expressed in distal regions in pre-osteoblasts, while bglap:GFP is only expressed in proximal regions in osteoblasts which do not express Runx2. Thus, bglap:GFP is activated in mature osteoblasts and the population does not include osteoblast precursor cells. These new data are found in Figure 2 —figure supplement 2B. In addition, we now also show by -RNAScope that bglap:GFP+ cells in segment -2 (the segment not affected by dedifferentiation) do not express the pre-osteoblast marker runx2a nor cyp26b1, which is upregulated in dedifferentiated osteoblasts. These additional data confirm that bglap:GFP specifically labels differentiated osteoblasts and does not include osteoblast precursors.

We have restructured the first paragraphs of the results to better describe our choice of markers and tools and describe the new data in the revised manuscript on page 6 as follows:

“(…) Using RNAScope in situ hybridization, we can now show that downregulation of bglap occurs in a graded manner and that entpd5 expression is similarly downregulated during dedifferentiation (Figure 1B, C). At 1 day post amputation (1 dpa), expression of entpd5 and bglap remains high in segment -2, but gradually decreases towards the amputation plane and is almost entirely absent from segment 0, with entpd5 downregulation being more pronounced (Figure 1B, C). While RNA expression of these genes is downregulated within hours after injury, GFP or Kaede fluorescent proteins (FPs) expressed in bglap or entpd5 reporter transgenic lines persist for up to three days, even though transgene transcription is shut down rapidly as well (Knopf et al., 2011). We can confirm these earlier findings using the more sensitive RNAScope in situs. In bglap:GFP transgenics at 2 dpa, gfp RNA and GFP protein colocalized to the same cells in segment -2, where osteoblasts do not dedifferentiate (Figure 1D). In contrast, in the distal segment -1 GFP protein was present, but barely any gfp transcript could be detected (Figure 1D). Thus, persistence of FPs in reporter lines can be used for short-term tracing of dedifferentiated osteoblasts (Figure 1E). At 1 dpa, bglap:GFP+ cells upregulated expression of the pre-osteoblast marker runx2a and of cyp26b1, an enzyme involved in retinoic acid signalling (Blum and Begemann, 2015), which regulates dedifferentiation (Figure 1 —figure supplement 1A, B). Both markers were exclusively upregulated in segment -1 and segment 0 at 1 dpa, but were absent in segment -2. Together, these data show that osteoblasts in segment -1 and segment 0 lose expression of mature markers and gain expression of dedifferentiation markers.”

3) Additional results are needed to support the claim that the complement system is required for injury–induced directed osteoblast migration including expression analyses of the central complement components C3 and C5 at the amputation site and showing whether c5ar1 and c3ar are solely expressed in osteoblasts, or rather broadly within tissue lining the hemirays.

Unfortunately, we lack the tools to detect the C3 and C5 precursor proteins or the mature cleavage products of the complement factors, which mediate the biological function of the cascade (e.g. antibodies against the zebrafish proteins / peptides). However, we analysed expression of the RNAs coding for the precursors of the complement factors c5 and the six zebrafish paralogs of c3 using qRT-PCR on liver, non-injured fins and fins at 6 hpa (samples derived from segment -1 plus segment 0). These new data can be found in Figure 5B. Compared to the expression levels in the liver, expression in non-injured fins could hardly be detected. Interestingly, c5 and c3a.5 levels were upregulated in injured fins, but compared to the expression in the liver still only slightly, e.g. c5 is about 17 Ct values (2 to the power of 17 = 130000 times) more highly expressed in the liver than in the injured fin. These results are consistent with the idea that the majority of complement factors that are activated after injury is derived from precursors that are expressed in the liver and are distributed via the circulation to the fin, as is considered standard for the complement system. Interestingly, however, local injury-induced expression in the fin might contribute precursor proteins as well.

While we had already shown that c5aR1 is expressed in osteoblasts, we have now added additional RNAscope in situ analysis for c5aR1 showing that the receptor is also expressed in other cell types in the fin (new data in Figure 5 —figure supplement 1A). We have also attempted RNAScope for c3aR1, c5a and c3a.1 in fins, but these turned out to not produce any specific stainings, thus the results of these experiments remained inconclusive and we have not included them in the manuscript. However, we have established fluorescent activated cell sorting from bglap:GFP transgenic fins, which gives us an additional tool to analyse to which extent expression is specific to osteoblasts. By qRT-PCR analysis we found that c5aR1 and c3aR are expressed in both GFP+ osteoblasts and other cells that are GFP– (these will mainly represent epidermis and fibroblasts, to a lesser extent endothelial and other cell types). These new data can be found in Figure 5 —figure supplement 1B.

While our qRT-PCR data and the c5aR1 RNAScope results show that the complement receptors are not specifically expressed in osteoblasts, we do not consider this result to be in conflict with our model that the complement system regulates osteoblast migration. Other cell types migrate after fin amputation as well, which is best described for epidermal cells (Chen et al., Dev Cell 2016, 10.1016/j.devcel.2016.02.017), but likely also occurs for fibroblasts (Poleo et al., DevDyn 2001, doi: 10.1002/dvdy.1152), and it is conceivable that the complement system plays a role in regulating these events as well.

Overall, our new data support our conclusion that the complement system is an important regulator of osteoblast migration in vivo, since the receptors are present in osteoblasts, while systemic and local expression can provide the precursors for injury-induced production of the activated factors that might act as guidance cues.

4) It is unclear whether each step of cell elongation and alignment, cell migration, cell dedifferentiation and regenerative response, is required for fin regeneration following amputation. In the absence of direct evidence for the requirement of migration or dedifferentiation for the overall success of fin regeneration, the limitations should be more clearly stated, and the title modified.

We have modified the title and abstract to avoid overstating the requirement of the particular responses to successful regeneration. Furthermore, we have stated the limitations of our study more clearly in the discussion.

The title now reads:

“Zebrafish fin regeneration involves generic and regeneration-specific osteoblast injury responses”

In the discussion we state the limitations on page 21 as follows:

“Unfortunately, currently existing tools to block dedifferentiation are either mosaic (activation of NF- κB signalling using the Cre-lox system) or cannot be targeted to osteoblasts alone (treatment with retinoic acid). Due to these limitations in our assays, we can currently not test what consequences specific, unmitigated perturbation of osteoblast dedifferentiation has for overall fin / bone regeneration. Conversely, the interventions presented here that specifically perturb osteoblast migration are limited as they act only transiently, that is they can severely delay, but not fully block migration. Furthermore, while interference with actomyosin dynamics reduces regenerative growth, we cannot distinguish whether this is caused by the inhibition of osteoblast migration or due to other more direct effects on cell proliferation and tissue growth. Thus, an unequivocal test of the importance of osteoblast migration for bone regeneration requires different tools.”

5) The word trauma should be clarified as the authors do not provide evidence for a non–healing phenotype in their trauma model.

We apologize if our use of the term trauma has caused confusion. We have simply used it interchangeably with “injury”. We have now removed all references to “trauma” in the text.

6) The conclusions referring to the relevance of the work to bone regeneration should be toned down and more focused on fin regeneration.

We have rephrased the conclusion to have it more centred on bone regeneration in the fin.

The relevant parts of the discussion on page 25 now read as follows:

“In conclusion, our findings support a model in which zebrafish fin bone regeneration involves both generic and regeneration-specific injury responses of osteoblasts. Morphology changes and directed migration towards the injury site as well as dedifferentiation represent generic responses that occur at all injuries even if they are not followed by regenerative bone formation. While migration and dedifferentiation can be uncoupled and are (at least partially) independently regulated, they appear to be triggered by signals that emanate from all bone injuries. In contrast, migration off the bone matrix into the bone defect, formation of a population of (pre-) osteoblasts and regenerative bone formation represent regeneration-specific responses that require additional signals that are only present at distal-facing injuries. The identification of molecular determinants of the generic vs regenerative responses will be an interesting avenue for future research.”

7) The manuscript is overall well written but could be improved to be less technical to help the readers, and the discussion should be reduced in length.

We have edited the manuscript, in particular we have tried to explain the background and logic of our setup and tools at the beginning of the results. We have also shortened the discussion.

Reviewer #1 (Recommendations for the authors):

To determine whether each step of cell elongation and alignment, cell migration, cell dedifferentiation and regenerative response, is required for fin regeneration following amputation the authors discuss p14 that other tools will be needed. However, later time points could be assessed in this study, not only to show the long–term effects of the treatment on healing but also to show how healing eventually resolves in the two models used here. Did the authors assess the long–term effects of inhibitor treatments on the regeneration process? Does blocking the complement system or actomyosin prevent regeneration? Are both generic (migration, dedifferentiation) and regenerative growth required for regeneration? Since there are limitations in the approaches used, the authors should modify the title and replace the word "requires". The use of the term "require" should be reconsidered also in the abstract and throughout the manuscript if the requirement for healing or a specific process is not demonstrated.

As a read-out for long-term effects of blocking the complement system or actomyosin, we have analysed two additional aspects. First, we analysed if the upregulation of Runx2 in segment 0 at 2 dpa is affected. We consider this another, but later readout for osteoblast dedifferentiation in addition to the bglap downregulation at 1 dpa, which we also show is not affected by actomyosin or complement inhibition (Figure 3F and Figure 5F). Neither actin nor complement interference had an effect on Runx2 upregulation. These new data can be found in Figure 3 —figure supplement 1B and Figure 5 —figure supplement 3B. These data confirm that these interventions do not block osteoblast dedifferentiation.

Second, we measured the length of the regenerate at 3 dpa as read-out for regenerative growth. Complement inhibition had no effect on regenerate length (new Figure 5 —figure supplement 3C). However, while osteoblast migration is severely impaired early after fin amputation upon complement inhibition, osteoblasts eventually do migrate to the amputation plane. We attempted to use higher doses of the drugs and the C3aR and C5aR1 inhibitors in combination, but these interventions turned out to be toxic and were thus inconclusive. Thus, the absence of regenerate growth defects upon complement inhibition cannot be interpreted to indicate that osteoblast migration is not required for fin regeneration.

In contrast, actomyosin inhibition did impair regenerative growth, (new data in Figure 3 —figure supplement 1C). Unfortunately, since actomyosin has essential roles in many cellular processes we cannot distinguish whether the impaired growth upon actomyosin inhibition is an indirect consequence of failed osteoblast migration or due to a direct role of actomyosin dynamics in cell proliferation and growth. Thus, the situation remains as we have explained in the discussion, that thorough tests of the individual requirements of the different osteoblast injury responses for overall regeneration will require additional tools. While this represents a limitation or our study, we still consider our main finding, that different injury responses can be uncoupled and are independently regulated, an important advance in our understanding of regeneration. We have revised the manuscript and replaced the word “require” with “involve”, e.g. in the title and abstract, which now read:

“Zebrafish fin regeneration involves generic and regeneration-specific osteoblast injury responses”

“Successful regeneration requires the coordinated execution of multiple cellular responses to injury. In amputated zebrafish fins, mature osteoblasts dedifferentiate, migrate towards the injury and form proliferative osteogenic blastema cells. We show that osteoblast migration is preceded by cell elongation and alignment along the proximodistal axis, which require actomyosin, but not microtubule turnover. Surprisingly, osteoblast dedifferentiation and migration can be uncoupled. Using pharmacological and genetic interventions, we found that NF-ĸB and retinoic acid signalling regulate dedifferentiation without affecting migration, while the complement system and actomyosin dynamics affect migration but not dedifferentiation. Furthermore, by removing bone at two locations within a fin ray, we established an injury model containing two injury sites. We found that osteoblasts dedifferentiate at and migrate towards both sites, while accumulation of osteogenic progenitor cells and regenerative bone formation only occur at the distal-facing injury. Together, these data indicate that osteoblast dedifferentiation and migration represent generic injury responses that are differentially regulated and can occur independently of each other and of regenerative growth. We conclude that successful bone regeneration appears to involve the coordinated execution of generic and regeneration-specific responses of osteoblasts to injury.”

Can the authors provide evidence of the source of C3 and C5, the central components of the complement system during fin regeneration? Figure 4 shows RNAscope in situ detection of c5aR1. Analysis of c3 and c5 using the same technique should be included.

Unfortunately, we lack the tools to detect the C3 and C5 precursor proteins or the mature cleavage products of the complement factors, which mediate the biological function of the cascade (e.g. antibodies against the zebrafish proteins / peptides). However, we analysed expression of the RNA coding for the precursors of the complement factors c5 and the six zebrafish paralogs of c3 using qRT-PCR on liver, non-injured fins and fins at 6 hpa (samples derived from segment -1 plus segment 0). These new data can be found in Figure 5B. Compared to the expression levels in the liver, expression in non-injured fins could hardly be detected. Interestingly, c5 and c3a.5 levels were upregulated in injured fins, but compared to the expression in the liver still only slightly, e.g. c5 is about 17 Ct values (2 to the power of 17 = 130000 times) more highly expressed in the liver than in the injured fin. These results are consistent with the idea that the majority of complement factors that are activated after injury is derived from precursors that are expressed in the liver and are distributed via the circulation to the fin, as is considered standard for the complement system. Interestingly, however, local, injury-induced expression might contribute precursor proteins as well.

Regarding the trauma model, it is unclear whether the gaps on each side of the remaining segment eventually heal. Does the gap on the proximal side of the remaining segment heal via regeneration occurring on the distal end of the next segment? The term "trauma" should be replaced by another term. Further, the term "trauma" appears to be used in the first paragraph of the discussion when referring to amputation and in other parts of the discussion when referring to bone injury.

In the hemiray injury model, the proximal side does indeed heal via regeneration form the distal end of the proximally located segment. We have added this information to the manuscript on page 16 as follows:

“Similarly, new bone matrix was formed at the distal side of the remaining proximal segment (arrowhead in Figure 6A and data not shown), but not at the proximal injury site.”

We apologize if our use of the term trauma has caused confusion. We have simply used it interchangeably with “injury”. We have now removed all references to “trauma” in the text.

The Discussion section is very long and there are many redundancies with the Results section. The main findings and conclusions could be stated more briefly, and the discussion points presented in a more concise way. As indicated in the public review, the last paragraph of the discussion should be focused on "fin regeneration" and avoid concluding on "bone regeneration" and "bone formation".

We have shortened the discussion to avoid redundancies, and have rephrased the conclusion to have it more centred on bone regeneration in the fin.

Reviewer #2 (Recommendations for the authors):

– Page 4, line 6: I am not sure if "exoskeletal" is by definition the right term in this context.

We thank the reviewer for the correction. We have rephrased the text on page 4 as follows:

“The fin skeleton consists of dermal (directly ossifying) skeletal elements, the fin rays or lepidotrichia, which make up the largest part of the fin, and endochondral parts close to the body.”

– What is the p–value in Figure 2A (left) between the width/length ratio of cytochalasin–treated osteoblasts from segments –1 and –2? It looks as if the cells of segment –1 are somewhat elongated even though they have been exposed to the drug. Could the authors please comment on this?

Upon cytochalasin treatment, osteoblasts in segment -1 are indeed elongated compared to segment -2 (p = 0.015, note that the data is now in Figure 3A). For the drug treatments affecting actomyosin, we had to carefully titrate drug concentrations such that the overall well-being of the fish was not affected. Thus, our hypothesis is that with the drug concentration we used, actin dynamics are impaired but not fully abolished. Therefore, one can still observe a cell shape change, but it is significantly decreased compared to non-treated fish.

– The red labeling of data points for the drug–treated conditions in Figure 2A, B and C might be difficult to be detected by color–blind readers. This should be adjusted accordingly.

As suggested by the reviewer, we have changed the color code.

– An image showing cyp26b1 RNAscope in situ hybridization on which the graphs of Figure 5H are based should be shown somewhere – maybe in the dedicated Excel sheet of raw data.

We have added a representative image of cyp26b1 RNAscope in situ hybridization (new data in Figure 6 —figure supplement 1B).

– Suggestion for a further experiment (not required for acceptance from my side!!!): I think it would be nice to learn whether C3a and C5a are sufficient to drive directed osteoblast migration along bony elements. To test that one could try to implant a bead loaded with recombinant C3a or C5a somewhere along a fin ray to see whether this ectopic signaling source attracts osteoblasts. Would this be doable? Recombinant complement proteins appear to be commercially available.

This is a great idea, and we have actually tried this by providing an ectopic source of the ligands next to the bony rays. However, it is technically very challenging to implant beads into the rather thin tissue between rays. Thus, we have not yet been able to derive conclusions from this experiment.

Reviewer #3 (Recommendations for the authors):

1. Title:

– I feel that the current title does not capture the essence of the authors' major findings.

In our study, we analyse several aspects (dedifferentiation, migration, proliferation) of osteoblast responses to amputation. We feel that the independence of these responses and the fact that dedifferentiation and migration can occur independently of regenerative bone growth are the main novel findings, which we tried to highlight in the title. However, since we agree with comments by several reviewers that we should be more cautious in proclaiming that all injury responses are “required”, we have replaced this term in the title with “involves”, the title now reads:

“Zebrafish fin regeneration involves generic and regeneration-specific osteoblast injury responses”

2. Abstract:

– The abstract is oversimplified and overstated, omitting important details. The authors should clearly state that they used a bglap:GFP strain to support their conclusions on osteoblast differentiation and migration.

As described above, we have several lines of additional evidence beyond the bglap:GFP line supporting our conclusions about osteoblast migration and its independence from dedifferentiation. We thus feel that we can use somewhat more general language in the abstract.

To avoid overstating the requirement of the particular responses to successful regeneration, we have modified the abstract. The last sentence now reads as follows:

“We conclude that successful bone regeneration appears to involve the coordinated execution of generic and regeneration-specific responses of osteoblasts to injury.”

3. Introduction:

– The authors make significant statements on osteoblast dedifferentiation. This is based on a naïve assumption that bglap:GFP is solely active in mature osteoblasts. However, it is clear from the authors' data that a substantial fraction of GFP+ cells are in proliferation (Figure 4 S1B). It seems that bglap:GFP is also expressed by osteoblast precursors. Specificity of bglap:GFP to mature osteoblasts is a major concern.

To address these comments, we have performed several additional experiments as described below. In addition, we would like to refer the reviewer to our previous papers, where we have analysed the process of osteoblast dedifferentiation (Knopf et al., Dev Cell 2011, doi: 10.1016/j.devcel.2011.04.014; Geurtzen et al., Development 2014, doi: 10.1242/dev.105817; Mishra et al. Dev Cell 2020, doi: 10.1016/j.devcel.2019.11.016). Using transgenic reporters and immunofluorescence we have shown in these previous papers that osteoblasts in the non-injured fin express Bglap but not the pre-osteoblast marker Runx2 (and are thus by our definition differentiated). We apologize if we failed to explain the logic of our approach in this manuscript, we have restructured the results to clarify these, as indicated below.

We have also performed the following additional experiments.

1) To confirm the specificity of the bglap:GFP line for mature osteoblasts, we have performed three experiments:

a) Immunofluorescence against Runx2 on 7 dpa regenerates, at a stage where blastema proliferation at the distal tip of the regenerate produces new osteoblast progenitors, while in more proximal (older) regions osteoblasts have already started to differentiate and new bone matrix has formed. We found that Runx2 is expressed in distal regions in pre-osteoblasts, while bglap:GFP is only expressed in proximal regions in osteoblasts which do not express Runx2. Thus, formation of new bony segment during regenerative growth, bglap:GFP is activated in mature osteoblasts and the population does not include osteoblast precursor cells. These new data are found in Figure 2 —figure supplement 2B.

b) We have refined and expanded our methods and are now able to determine the expression patterns of markers of the osteoblast differentiation status with single cell resolution using RNAScope in situ hybridization. Using this, we can now show that at 1 day post amputation, in segment -2 of the fin stump, which represents a segment equivalent to the non-injured state, since no dedifferentiation occurs here, bglap:GFP+ cells do not express endogenous runx2a. These new data are found in Figure 1 —figure supplement 1A.

c) Using RNAScope, we can show that cyp26b1, a gene associated with dedifferentiated osteoblasts, is likewise not detected in bglap:GFP+ cells in segment -2 at 1 dpa (new data in Figure 1 —figure supplement 1B).

Together, these data confirm that the bglap:GFP line is specific for differentiated osteoblasts, and does not label osteoblast progenitors.

See the response to issue 2 below for how we describe these new data in the revised version of the manuscript.

2) Regarding the proliferation of bglap:GFP osteoblasts: In the experiment the reviewer refers to (now Figure 5 —figure supplement 3A), we make use of the persistence of the GFP protein in the bglap:GFP line to detect dedifferentiated osteoblasts. Thus, at the time of analysis, when these GFP+ cells proliferate, they are not differentiated anymore. We can show this as follows:

Although bglap expression is downregulated during osteoblast dedifferentiation and thus also GFP levels eventually drop in the transgenic line, we can nevertheless use this line to trace osteoblasts, since GFP protein persists for up to three days in cells that shut down endogenous bglap and also bglap:GFP transgene transcription. While we have already shown this previously (Knopf et al., Dev Cell 2011, doi: 10.1016/j.devcel.2011.04.014; Geurtzen et al., Development 2014, doi: 10.1242/dev.105817; Mishra et al. Dev Cell 2020, doi: 10.1016/j.devcel.2019.11.016), we have now also used RNAScope to confirm this. We analysed the expression of GFP on protein and RNA level in the bglap:GFP line. In bglap:GFP fish, in a mature segment in non-injured fins the regions close to the joints are devoid of cells expressing GFP (Figure 1G). Yet after amputation, we observe GFP+ cells in this distal part of segment -1 (Figure 1G, D). RNAscope in situ shows that these GFP+ cells are negative for gfp RNA (new data in Figure 1D). Thus, the observed fluorescence is due to the persistence of the GFP protein and not due to a potential upregulation of the transgene (Figure 1E).

Importantly, we have now also added data describing the proliferative state of bglap:GFP+ osteoblasts. First, in the non-injured fin, bglap:GFP+ cells are non-proliferative (new data in Figure 5 —figure supplement 2B). After amputation, proliferation can be detected in GFP+ cells at 2 dpa (Figure 5 —figure supplement 2B), and proliferation is restricted to segment -1 and segment 0 (new data in Figure 5 —figure supplement 2C). As we show in Figure 1B, at 2 dpa, dedifferentiation as defined by bglap downregulation is not complete in segment -1, rather here a mixture of cells with different bglap levels are found. We have thus combined EdU labelling with RNAscope against bglap in segment -1 to analyse to which extent bglap and EdU anticorrelate. These data show that EdU is hardly ever incorporated into cells expressing high levels of bglap, while the majority of the proliferating osteoblasts are dedifferentiated, as they express only low levels of bglap (new data in Figure 5 —figure supplement 2D). Together, these data show that mature osteoblasts are non-proliferative, and upon amputation, when they are dedifferentiated, they become proliferative. Thus, the absence of proliferation in bglap:GFP+ cells in the non-injured fin adds to the evidence that this line is specific for mature osteoblasts, but due to the persistence of the GFP protein it can be used to analyse dedifferentiated osteoblasts.

These data are described on page 14 of the manuscript as follows:

“In the non-injured fin, bglap:GFP+ osteoblasts are non-proliferative, but upon amputation osteoblasts proliferate at 2 dpa (Figure 5 —figure supplement 2A, B). Proliferation is restricted to segment -1 and segment 0 (Figure 5 —figure supplement 2C), and RNAscope in situ analysis of bglap expression revealed that the majority of EdU+ osteoblasts have strongly downregulated bglap (Figure 5 —figure supplement 2D). Inhibition of C5aR1 with PMX205 had no effect on osteoblast proliferation in segment -1 at 2 dpa (Figure 5 —figure supplement 3A). Furthermore, upregulation of Runx2 was not changed by PMX205 treatment (Figure 5 —figure supplement 3B), and regenerative growth was not affected in fish treated with either W54011, PMX205 or SB290157 (Figure 5 —figure supplement Figure 3C). We conclude that the complement system specifically regulates injury-induced osteoblast migration, but not osteoblast dedifferentiation or proliferation in zebrafish.”

3) To support our conclusion that osteoblasts migrate, we performed time-lapse imaging using a transgenic line expressing the photoconvertible protein kaede in osteoblasts (entpd5:kaede). Local photoconversion of only the proximal half of a segment allowed us to trace these photoconverted osteoblasts. This revealed that converted cells appear in the distal part of the segment within 1 dpa, which can only be explained by relocation of the cells. These new data can be found in Figure 1F and they are described on page 7 of the revised manuscript as follows:

“To trace osteoblasts, we used the transgenic line entpd5:kaede (Geurtzen et al., 2014), in which Kaede fluorescence can be converted from green to red by UV light (Ando et al., 2002). We photoconverted osteoblasts in the proximal half of segment -1, while osteoblasts in the distal half remained green (Figure 1F). At 1 dpa, red osteoblasts were found in the distal half (Figure 1F), showing that photoconverted osteoblasts had relocated distally.”

– Page 5, Line 5: Osteoblasts do not migrate to the resorbed sites. Osteoblast precursors (or pre–osteoblasts) do. Thiel 2018 review article does not appear to be particularly supportive.

We thank the reviewer for the correction. We have corrected it and added another reference:

“For bone formation, osteoblast precursors migrate to the resorbed sites (Dirckx, Van Hul and Maes, 2013)”

4. Results:

– Page 6, Line 14: Perhaps the authors should explain in more detail about the kaedeGreen to Red system, and why they needed this approach.

We have expanded the description of the photoconvertible kaede system. It is first introduced now on page 7 (describing a new experiment), where it reads as follows:

“To trace osteoblasts, we used the transgenic line entpd5:kaede (Geurtzen et al., 2014), in which Kaede fluorescence can be converted from green to red by UV light (Ando et al., 2002).”

In the experiment the reviewer is referring to, we used a double transgenic line expressing both GFP and kaede in osteoblasts (Video 1, Figure 2 —figure supplement 1A). We deliberately only partly converted kaedeGreen to kaedeRed, which resulted in different hues for each osteoblast. This distinct colouring facilitates identification of individual cells during repeated live imaging using a confocal microscope. Due to the curved surface of a hemiray and to counteract potential movements along the z-axis of the fin during image acquisition, several z-slides had to be recorded at each timepoint. As a consequence, there is a considerable time of recording, increasing photobleaching over time. Having two fluorescent proteins being expressed in the cells gave a higher intensity signal in the beginning, allowing lower laser power and exposure time, and thus reduced photobleaching. The description now reads as follows (page 8):

“To observe osteoblast behaviour at single cell resolution in live fish after fin amputation, we generated double transgenic fish expressing bglap:GFP (Knopf et al., 2011)and entpd5:kaede in mature osteoblasts. Expression of two fluorescent proteins increased signal for repeated imaging, and partial photoconversion of kaedeGreen into kaedeRed resulted in a different colouring for each osteoblast, which facilitated tracking of morphology changes at single cell resolution.”

– Page 7, Line 13–16: I do not understand why the authors interpret these data as directed active migration behavior towards the injury. What they showed here is merely cell shape changes.

The formation of protrusions orientated in the direction of (nascent) movement is an essential step during cell migration. Our data show that mature osteoblasts in the non-injured fin are roundish without such directed protrusion. In contrast, after amputation we see an elongation of osteoblasts in segment -1 in the direction of migration. We agree that such protrusions on their own would not be sufficient evidence of active migration. Yet, in combination with our other evidence (relocation of cells shown by bglap:GFP and entpd5:Kaede tracing via repeated imaging, live videos showing movement of osteoblasts relative to their surroundings, see the response to “Weaknesses”, issue 3 above), we interpret this morphology changes as being consistent with acquisition of active migratory behaviour. To better elucidate the relation between cell shape changes and migration, we have re-arranged the order in which we present our data. We now show the relocation / migration of osteoblasts first (Figure 1D – I), and then the cell shape changes (Figure 2A-C, Figure 2 —figure supplement 1). This highlights that only in the segments where osteoblasts migrate, they elongate. Furthermore, we have rephrased the sentence. It now reads as follows (page 9):

“We interpret the elongation and orientation along the proximodistal axis and the formation of long-lived protrusions along this axis as events that prime osteoblasts for directed active migration towards the injury.”

– Page 7, Line 19: This data could be alternatively interpreted as the upregulation of bglap:GFP by invading cells (not previously bglap+), rather than the invasion of bglap:GFP+ cells. It would be essential to determine the half–life of GFP to assume cell invasion. The authors would need a more long–lasting histone–bound GFP reporter strain to conclusively define cell invasion.

Please see our response above. There we describe new experiments where we detect gfp RNA and GFP protein to show that appearance of GFP+ cells is indeed due to invasion, not upregulation of transgene transcription. See new data in Figure 1D, E.

– Page 8, Line 3–16: This paragraph is very difficult to read because of long sentences. They should simplify and rephrase the sentences.

We have rephrased this paragraph and shortened the sentences. It now reads as follows (page 9):

“Fins grow by the addition of new segments distally, and in the distal-most, youngest segment, bglap:GFP is not expressed (Knopf et al., 2011). Similarly, in the regenerating fins increasing numbers of GFP+ cells can be detected in more proximally located, older segments, reflecting the progressive differentiation of osteoblasts with time (Figure 2 —figure supplement 2A). In less mature segments, the pre-osteoblast marker Runx2 can be detected, but its expression does not overlap with bglap:GFP expression in mature osteoblasts (Figure 2 —figure supplement 2B). Importantly, in newly regenerated segments that start to upregulate bglap expression, all bglap:GFP+ cells appear in the centre of the segments; we could not observe GFP+ cells within joints (Figure 2 —figure supplement 2A). This suggests that bglap:GFP+ osteoblasts from older segments do not migrate into less mature segments during formation of new segments in the course of fin growth. Rather, the mature osteoblast population in a segment is formed via differentiation of osteoblasts at the position within the segment where they were formed during segment formation. In contrast, within 2 days after fin amputation, bglap:GFP+ cells appeared in the fin stump within the joint between segment -1 and segment 0 (Figure 2D, yellow arrowhead), indicating that GFP+ osteoblasts from segment -1 crossed the joint during their migration towards the amputation plane. Thus, migration of mature osteoblasts observed after fin amputation or bone fracture (Geurtzen et al., 2014) appears to represent a specific early response to injury, while addition of new bony segments at the distal tip of growing fins during ontogeny or regeneration does not involve migration of differentiated osteoblasts.”

– Page 9, Line 1: The authors should move the bglap RNAscope in situ data to the main figure, as this is an important piece of information.

We agree that these data were somewhat hidden in the Supplement, we have now moved them to the re-designed Figure 1B.

– Page 9, Line 6: I don't understand why reduced bglap mRNA can be interpreted as osteoblast dedifferentiation.

We define dedifferentiation as the reversion of a mature cell into an undifferentiated progenitor-like status. This involves the following characteristics: (1) the expression of markers of the differentiated state are downregulated; (2) early lineage markers are re-expressed; (3) the cells become proliferative; and (4) they have the ability to re-differentiate into mature cells. Based in this definition, the downregulation of an osteoblast-specific marker can be used as a read-out for osteoblast dedifferentiation. Bglap is an established marker for mature osteoblasts (Kaneto et al., 2016 doi.org/10.1186/s12881-016-0301-7¸ Yoshioka et al., 2021 doi: 10.1002/jbm4.10496; Kannan et al., 2020 doi: 10.1242/bio.053280; Sojan et al., 2022 doi.org/10.3389/fnut.2022.868805; Valenti et al., 2020 doi.org/10.3390/cells9081911). While we use downregulation of bglap expression as our main read-out for osteoblast dedifferentiation in our experimental interventions (actomyosin inhibition, retinoic acid treatment, complement inhibition), we have expanded our methods to characterize osteoblast dedifferentiation, and have re-arranged our manuscript to show these data in the beginning of the results.

Already in the previous version of the manuscript we have shown that endogenous bglap is strongly expressed in segment -2, (the segment that does not respond to fin amputation and thus represents the non-injured state), while it is downregulated in a graded manner in segment -1 and segment 0 (the segments where dedifferentiation happens). We have now moved this data to the re-designed Figure 1B. In addition to bglap, we can now show that entpd5, a gene required for bone mineralization, is strongly expressed in osteoblasts of segment -2, while it is massively downregulated in segment -1 and segment 0. These new data can be found in Figure 1C. Thus, entpd5 is another differentiation marker whose loss characterizes osteoblast dedifferentiation. Importantly, we can confirm by RNAScope that the pre-osteoblast marker runx2a is absent in mature segments but is upregulated in segment 0 and segment -1 at 1 dpa (new data in Figure 1 —figure supplement 1A). Similarly, cyp26b1, an enzyme shown to regulate dedifferentiation, is upregulated in segment 0 and segment -1, but not expressed in segment -2. (new data in Figure 1 —figure supplement 1B). Furthermore, we have repeated all experiments where we have previously quantified dedifferentiation upon experimental interventions using downregulation of bglap:GFP (actomyosin inhibition, retinoic acid treatment, complement inhibition). We now can fully confirm the previous conclusions using the more rigorous quantification of dedifferentiation using RNAScope analysis of endogenous bglap levels. We have replaced all bglap:GFP data with the new bglap RNAScope data. These new data are found in Figure 3F, Figure 3 —figure supplement 1A, Figure 4B and Figure 5F.

Overall, we support our conclusion that osteoblasts dedifferentiate by the loss of the two differentiation markers bglap and entpd5, the upregulation of the pre-osteoblast marker runx2a and the dedifferentiation-associated gene cyp26b1, and the fact that osteoblasts become proliferative. We hope that the reviewer considers this sufficient evidence.

In mammals, the available literature relatively convincingly concludes that NF-κB signaling negatively regulates osteoblast differentiation (Yao et al., 2014, doi: 10.1002/jbmr.2108; Swarnkar et al., 2014 doi.org/10.1371/journal.pone.0091421, Chang et al., 2009, doi.org/10.1038/nm.1954). Yet in zebrafish osteoblasts, we have previously shown that NF-κB signaling is active in mature osteoblasts and needs to be downregulated for dedifferentiation to occur (Mishra et al., 2020, 10.1016/j.devcel.2019.11.016). Importantly, in our previous work we showed that at least during fin regeneration, NF-κB signalling is not involved in osteoblast differentiation (Mishra et al., 2020, 10.1016/j.devcel.2019.11.016). Specifically, osteoblasts in which Nf-kappaB signaling is enhanced or inhibited differentiate completely normally during the later stages of fin regeneration in the fin regenerate. Hence, our findings with the Nf-kappaB intervention studies done in this manuscript, where we look at osteoblasts in the stump within 1 dpa, cannot be explained by them affecting osteoblast differentiation.

For retinoic acid signalling, multiple roles in bone development and repair have been described in mammals. For zebrafish osteoblasts, it was shown that during the outgrowth phase of bone regeneration, retinoic acid negatively regulates osteoblast differentiation in the blastema (Blum & Begemann, 2015, 10.1242/dev.120204). Yet importantly, it also negatively controls the dedifferentiation of osteoblasts in the stump right after amputation (Blum & Begemann, 2015, 10.1242/dev.120204). Thus, the effect we observe at the early timepoints we analyse in our intervention studies (retinoic acid treatment) are due to the effect on osteoblast dedifferentiation.

We have added a short definition of dedifferentiation to the Results section (page 6). There it reads as follows:

“We have previously shown that osteoblasts dedifferentiate in response to fin amputation, that is they revert from a mature, non-proliferative state into an undifferentiated progenitor-like state, which includes loss of bglap expression and upregulation of the pre-osteoblast marker runx2 (Knopf et al., 2011; Geurtzen et al., 2014).”

In addition, we have restructured the results to describe our use of tools and the new data on page 6 of the revised manuscript as follows:

“Using RNAScope in situ hybridization, we can now show that downregulation of bglap occurs in a graded manner and that entpd5 expression is similarly downregulated during dedifferentiation (Figure 1B, C). At 1 day post amputation (1 dpa), expression of entpd5 and bglap remains high in segment -2, but gradually decreases towards the amputation plane and is almost entirely absent from segment 0, with entpd5 downregulation being more pronounced (Figure 1B, C). While RNA expression of these genes is downregulated within hours after injury, GFP or Kaede fluorescent proteins (FPs) expressed in bglap or entpd5 reporter transgenic lines persist for up to three days, even though transgene transcription is shut down rapidly as well (Knopf et al., 2011). We can confirm these earlier findings using the more sensitive RNAScope in situs. In bglap:GFP transgenics at 2 dpa, gfp RNA and GFP protein colocalized to the same cells in segment -2, where osteoblasts do not dedifferentiate (Figure 1D). In contrast, in the distal segment -1 GFP protein was present, but barely any gfp transcript could be detected (Figure 1D). Thus, persistence of FPs in reporter lines can be used for short-term tracing of dedifferentiated osteoblasts (Figure 1E). At 1 dpa, bglap:GFP+ cells upregulated expression of the pre-osteoblast marker runx2a and of cyp26b1, an enzyme involved in retinoic acid signalling (Blum and Begemann, 2015), which regulates dedifferentiation (Figure 1 —figure supplement 1A, B). Both markers were exclusively upregulated in segment -1 and segment 0 at 1 dpa, but were absent in segment -2. Together, these data show that osteoblasts in segment -1 and segment 0 lose expression of mature markers and gain expression of dedifferentiation markers.”

– Overall, the authors would need live–cell imaging and tracking to make these conclusions.

We have the following evidence for osteoblast migration:

1) bglap:GFP+ cells relocate from the centre of segments towards the amputation plane (after fin amputations) or towards both injuries in the hemiray model. In this revised manuscript we show that transgene expression is not upregulated in these regions, but that GFP fluorescence there must be due to relocation of cells in which GFP protein persists (new data in Figure 1D, E; see also response to “Weaknesses, issue 1” above)

2) Using the entpd5:kaede transgenic line, which is expressed in mature osteoblasts throughout segments, we have photoconverted only the proximal half of a segment, which allowed us to trace these photoconverted osteoblasts. This revealed that converted cells appear in the distal part of the segment within 1 dpa, which can only be explained by relocation of the cells. These new data can be found in Figure 1F.

3) Already in the previous version of the manuscript, we have performed live imaging to track single cell behaviour. Using double transgenic fish expressing both GFP and kaede in osteoblasts, we deliberately only partly converted kaedeGreen to kaedeRed, which resulted in different hues for each osteoblast. This distinct colouring facilitates observing single cells. Video 1 shows the directed movement of cell bodies relative to their surroundings within 2 hours (see also Figure 2 —figure supplement 1A).

4) Osteoblasts display the typical cell shape changes associated with active migration (elongation along the axis of migration, extension of dynamic protrusions), data in Figure 2.

Together, we think these are convincing data supporting the conclusion that osteoblasts actively migrate.

– Page 10, Line 7: If the authors think that a bglap:GFP strain can be used as live reporters for osteoblast dedifferentiation, that would significantly compromise the scientific rigor of the study, as it would not support any conclusions regarding cell migration and invasion.

Please see the response to “Weaknesses”, issue 1 above. As we have laid out there, bglap:GFP can be used for both short-term tracing of cells that used to be differentiated due to persistence of GFP protein after transgene shutdown during dedifferentiation, and as readout for dedifferentiation, since GFP levels eventually do drop as well. However, we agree that its use as a readout for dedifferentiation is complicated and not straightforward. Thus, we have repeated all experiments where we have previously quantified dedifferentiation upon experimental interventions using downregulation of bglap:GFP (actomyosin inhibition, retinoic acid treatment, complement inhibition). We now can fully confirm the previous conclusions using the more rigorous quantification of dedifferentiation using RNAScope analysis of endogenous bglap levels. These new data are found in Figure 3F, Figure 3 —figure supplement 1A, Figures 4B, Figure 5F.

– Page 10, Line 20: Osteoblast migration should not be mixed up with cell shape changes.

We agree that cell shape changes and relocation of cells relative to their surroundings are strictly not the same process. Yet our data support the idea that both are events associated with the same process, namely osteoblast migration. This recapitulating sentence refers to the data we presented regarding the effect of actomyosin and microtubuli drugs. For both interventions we have assayed osteoblast relocation (of bglap:GFP+ cells from the centre towards the distal end of the segment) and cell shape changes. We thus think that both assays together allow us to describe these data as affecting osteoblast “migration”. This sentence is now on page 12, line 1.

– Page 11, Line 12: The authors did not show data for osteoblast differentiation/dedifferentiation in their genetic intervention studies.

The genetic intervention was used to manipulate NF-κB signalling. There are established tools available to genetically interfere with NF-κB signalling, and we used such transgenic tools in our assays. NF-κB signalling can be inhibited by overexpression of the IkB super repressor (IkBSR) (Van Antwerp et al., 1996 doi: 10.1126/science.274.5288.787; Karra et al., 2015 DOI: 10.1073/pnas.1511209112.), while overexpression of a constitutively active IkB kinase (IKKca) results in overactivation of the pathway (Mishra et al., 2020 10.1016/j.devcel.2019.11.016). To specifically manipulate the pathway in osteoblasts, we employ the Cre-Lox system, using the established inducible line osx:CreER (Knopf et al., 2011, doi: 10.1016/j.devcel.2011.04.014). We have previously shown that such osteoblast-specific overexpression of IkBSR enhanced osteoblast dedifferentiation, while overexpression of IKKca impaired dedifferentiation (Mishra et al., 2020 10.1016/j.devcel.2019.11.016). Due to technical reasons, the assay to test the effect of the genetic interventions on dedifferentiation cannot be combined with the read-out for osteoblast cell shape. Yet, since we have shown the effect of these tools on dedifferentiation in our previous paper, we did not repeat these assays for this manuscript.

– Page 11, Line 19: An alternative explanation is that retinoic acid and NF–kB signaling simply promote osteoblast differentiation, without anything to do with dedifferentiation.

In mammals, the available literature relatively convincingly concludes that NF-κB signaling negatively regulates osteoblast differentiation (Yao et al., 2014, doi: 10.1002/jbmr.2108; Swarnkar et al., 2014 doi.org/10.1371/journal.pone.0091421, Chang et al., 2009, doi.org/10.1038/nm.1954). Yet in zebrafish osteoblasts, we have previously shown that NF-κB signaling is active in mature osteoblasts and needs to be downregulated for dedifferentiation to occur (Mishra et al., 2020, 10.1016/j.devcel.2019.11.016). Importantly, in our previous work we showed that at least during fin regeneration, NF-κB signalling is not involved in osteoblast differentiation (Mishra et al., 2020, 10.1016/j.devcel.2019.11.016). Specifically, osteoblasts in which Nf-kappaB signaling is enhanced or inhibited differentiate completely normally during the later stages of fin regeneration in the fin regenerate. Hence, our findings with the Nf-kappaB intervention studies done in this manuscript, where we look at osteoblasts in the stump within 1 dpa cannot be explained by them affecting osteoblast differentiation.

For retinoic acid signalling, multiple roles in bone development and repair have been described in mammals. For zebrafish osteoblasts, it was shown that during the outgrowth phase of bone regeneration, retinoic acid negatively regulates osteoblast differentiation in the blastema (Blum & Begemann, 2015, 10.1242/dev.120204). Yet importantly, it also negatively controls the dedifferentiation of osteoblasts in the stump right after amputation (Blum & Begemann, 2015, 10.1242/dev.120204). Thus, the effect we observe at the early timepoints we analyse in our intervention studies (retinoic acid treatment) are due to the effect on osteoblast dedifferentiation

– Page 12, Line 1–2: These conclusions are stretched and not supported by experimental data. The argument for the decoupling of dedifferentiation and migration is not well developed.

We politely disagree with the reviewer as we do think that our data support the hypothesis that dedifferentiation and migration are not coupled in the sense that they are necessary consequences that are triggered by a single event, namely injury, and thus always occur together. We base our hypothesis on the following observations: we define molecular interventions in which (1) osteoblast elongation and migration are impaired, yet dedifferentiation is not affected (interference with complement and actomyosin systems, Figure 3), and (2) interventions where dedifferentiation is inhibited, yet osteoblasts still elongate and migrate (Nf-kappaB and retinoic acid signaling, Figure 4). If these two aspects were interdependent, one would expect the mitigation of one inevitably affecting the strength of the other. Yet we do not see such an effect but instead one can be diminished without the other being changed. Therefore, we conclude that they are independently regulated. However, we have rephrased the sentence to attenuate our conclusions about the independency of dedifferentiation and migration. It now reads as follows:

“Together, these data suggest that osteoblast dedifferentiation and migration are independently regulated, as one response can be impaired without affecting the other. Furthermore, they indicate that osteoblast dedifferentiation is not a prerequisite for migration.”

– Page 13, Line 8: As many as 20% of bglap:GFP+ cells are EdU+ therefore in proliferation. How would the authors explain this?

Please see the response to “Weaknesses”, issue 1, our point 2 above. In brief, the GFP+ cells analysed here are already dedifferentiated, yet GFP protein persists.

5. Discussion:

– The authors should clearly indicate their definition of osteoblasts in this study (i.e. bglap:GFP+ cells).

We state our definition of differentiated osteoblasts in the introduction as follows (page 4):

“…osteoblasts close to the amputation plane dedifferentiate, that is they downregulate the expression of the mature osteoblast marker bglap, upregulate pre-osteoblast markers like runx2 and become proliferative…

And in the results as follows (page 6):

“We have previously shown that osteoblasts dedifferentiate in response to fin amputation, that is they revert from a mature, non-proliferative state into an undifferentiated progenitor-like state, which includes loss of bglap expression and upregulation of the pre-osteoblast marker runx2.”

And in the discussion again as follows (page 20):

“In addition, osteoblasts dedifferentiate, that is they downregulate expression of differentiation markers like bglap and initiate expression of the pre-osteoblast marker runx2.”

– Osteoblast dedifferentiation should be rephrased as reduced bglap expression.

We agree that the term “dedifferentiation” is unfortunately loosely defined in the literature, in particular in relation to the question whether a process described as “dedifferentiation” would include a fate reversal of cells. However, since most instances of naturally occurring dedifferentiation during unperturbed regenerative events involve reversal of differentiated cells to a proliferative, progenitor-like state in which the cells remain lineage restricted, we favour a definition as follows:

Dedifferentiation is the reversion of a mature cell into an undifferentiated progenitor-like status. This involves the following characteristics: (1) the expression of differentiation markers is downregulated; (2) early lineage markers are re-expressed; (3) the cells become proliferative; and (4) they have the ability to re-differentiate into mature cells.

Already in the previous version of the manuscript, in the first mention of dedifferentiation in the discussion, we define dedifferentiation as loss of bglap and gain of the pre-osteoblast marker runx2. It reads as follows (page 20):

“In addition, osteoblasts dedifferentiate, that is they downregulate expression of differentiation markers like bglap and initiate expression of the pre-osteoblast marker runx2.”

– Page 18, Line 19: Osteoblast differentiation, instead of osteoblast dedifferentiation.

We have to respectfully disagree with the reviewer, as we do indeed mean dedifferentiation. As explained in detail in our answer to the public review, point 2, in the zebrafish fin, after amputation both pathways (NF-κB and retinoic acid) regulate osteoblast dedifferentiation, which was shown previously by us and other labs. This sentence is now page 20:

“Retinoic acid and NF-κB signalling pathways regulate only osteoblast dedifferentiation, but not cell shape changes and migration, while the complement and actomyosin systems are required for osteoblast cell shape changes and migration, but not for dedifferentiation.”

https://doi.org/10.7554/eLife.77614.sa2

Article and author information

Author details

  1. Ivonne Sehring

    Institute of Biochemistry and Molecular Biology, University of Ulm, Ulm, Germany
    Contribution
    Conceptualization, Formal analysis, Funding acquisition, Investigation, Methodology, Validation, Visualization, Writing – original draft
    For correspondence
    Ivonne.sehring@uni-ulm.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-7812-0278
  2. Hossein Falah Mohammadi

    Institute of Biochemistry and Molecular Biology, University of Ulm, Ulm, Germany
    Contribution
    Formal analysis, Investigation
    Competing interests
    No competing interests declared
  3. Melanie Haffner-Luntzer

    Institute of Orthopaedic Research and Biomechanics, University Hospital Ulm, Ulm, Germany
    Contribution
    Resources, Writing - review and editing
    Competing interests
    No competing interests declared
  4. Anita Ignatius

    Institute of Orthopaedic Research and Biomechanics, University Hospital Ulm, Ulm, Germany
    Contribution
    Resources, Writing - review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-4782-1979
  5. Markus Huber-Lang

    Institute of Clinical and Experimental Trauma-Immunology (ITI), University Hospital Ulm, Ulm, Germany
    Contribution
    Resources, Writing - review and editing
    Competing interests
    No competing interests declared
  6. Gilbert Weidinger

    Institute of Biochemistry and Molecular Biology, University of Ulm, Ulm, Germany
    Contribution
    Conceptualization, Funding acquisition, Methodology, Project administration, Resources, Supervision, Writing – original draft
    For correspondence
    gilbert.weidinger@uni-ulm.de
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-3599-6760

Funding

Deutsche Forschungsgemeinschaft (Project-ID 251293561 - SFB 1149)

  • Gilbert Weidinger

Deutsche Forschungsgemeinschaft (project ID 316249678 - SFB 1279)

  • Gilbert Weidinger

Deutsche Forschungsgemeinschaft (project ID 450627322 - SFB 1506)

  • Gilbert Weidinger

Medical Faculty, Ulm University (Hertha-Nathorff-Program)

  • Ivonne Sehring

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

Acknowledgements

We thank Doris Weber and Marion Gradl for their contributions to fish care and the core facility 'Confocal and multiphoton imaging' of the Medical faculty of Ulm University for help with imaging. This work was funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) Project-ID 251293561 – SFB 1149, project ID 316249678 – SFB 1279, and project ID 450627322 – SFB 1506. Ivonne Sehring was funded by the Hertha-Nathorff-Program of the Medical Faculty of Ulm University.

Ethics

All procedures involving animals adhered to EU directive 2010/63/EU on the protection of animals used for scientific purposes, and were approved by the state of Baden-Württemberg (Project numbers 1193 and 1494) and by local animal experiment committees. Fish of both sexes were used. Housing and husbandry followed the recommendations of the Federation of European Laboratory Animal Science Associations (FELASA) and the European Society for Fish Models in Biology and Medicine (EUFishBioMed) (Aleström et al., 2020).

Senior Editor

  1. Didier YR Stainier, Max Planck Institute for Heart and Lung Research, Germany

Reviewing Editor

  1. Céline Colnot, INSERM U955, UNIV PARIS EST CRETEIL, France

Reviewer

  1. Céline Colnot, INSERM U955, UNIV PARIS EST CRETEIL, France

Publication history

  1. Received: February 4, 2022
  2. Preprint posted: February 22, 2022 (view preprint)
  3. Accepted: June 23, 2022
  4. Accepted Manuscript published: June 24, 2022 (version 1)
  5. Version of Record published: July 6, 2022 (version 2)

Copyright

© 2022, Sehring et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Ivonne Sehring
  2. Hossein Falah Mohammadi
  3. Melanie Haffner-Luntzer
  4. Anita Ignatius
  5. Markus Huber-Lang
  6. Gilbert Weidinger
(2022)
Zebrafish fin regeneration involves generic and regeneration-specific osteoblast injury responses
eLife 11:e77614.
https://doi.org/10.7554/eLife.77614
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    Ge Gao, Shuyu Guo ... Gang Peng
    Research Article Updated

    Unbiased genetic screens implicated a number of uncharacterized genes in hearing loss, suggesting some biological processes required for auditory function remain unexplored. Loss of Kiaa1024L/Minar2, a previously understudied gene, caused deafness in mice, but how it functioned in the hearing was unclear. Here, we show that disruption of kiaa1024L/minar2 causes hearing loss in the zebrafish. Defects in mechanotransduction, longer and thinner hair bundles, and enlarged apical lysosomes in hair cells are observed in the kiaa1024L/minar2 mutant. In cultured cells, Kiaa1024L/Minar2 is mainly localized to lysosomes, and its overexpression recruits cholesterol and increases cholesterol labeling. Strikingly, cholesterol is highly enriched in the hair bundle membrane, and loss of kiaa1024L/minar2 reduces cholesterol localization to the hair bundles. Lowering cholesterol levels aggravates, while increasing cholesterol levels rescues the hair cell defects in the kiaa1024L/minar2 mutant. Therefore, cholesterol plays an essential role in hair bundles, and Kiaa1024L/Minar2 regulates cholesterol distribution and homeostasis to ensure normal hearing.