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Dynamics of caspase-3-mediated apoptosis during spinal cord regeneration in the teleost fish, Apteronotus leptorhynchus
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Dynamics of caspase-3-mediated apoptosis during spinal cord regeneration in the teleost fish, Apteronotus leptorhynchus Ruxandra F. Sîrbulescu a , Günther K.H. Zupanc a,b,⁎ a
School of Engineering and Science, Jacobs University Bremen, P.O. BOX 750 561, 28725 Bremen, Germany Department of Biology, Northeastern University, 360 Huntington Avenue, Boston, MA 02115, USA
b
A R T I C LE I N FO
AB S T R A C T
Article history:
In contrast to mammals, adult teleost fish exhibit a vast potential for central nervous
Accepted 17 September 2009
system regeneration after injury. Among other mechanisms, this capacity is mediated by
Available online 24 September 2009
replacement of cells lost to injury by new neurons and glia. Here, we examined the spatio-
Keywords:
differentiation of new cells, during this cell replacement phase. As an experimental
temporal dynamics of apoptosis, and its relationship to the generation and the Spinal cord regeneration
paradigm, caudal transection of the spinal cord in the teleost fish Apteronotus leptorhynchus
Apoptosis
was used. During the cell replacement phase, there was a rather constant percentage of new
Cell proliferation
cells (identified by incorporation of 5-bromo-2′-deoxyuridine into newly synthesized DNA)
Adult neurogenesis
that underwent apoptosis (identified by anti-active caspase-3 immunolabeling). Many of
Caspase-3
these cells were also immunopositive for the marker proteins Hu C/D or glial fibrillary acidic protein, indicating that a large portion of cells undergo apoptosis after differentiation into neurons or glia, respectively. The spatial distribution of apoptotic cells was uneven, displaying a radial peak in the mid parenchymal regions and a longitudinal peak at the site of the initial spinal transection. The latter persisted for over 100 days post-injury, indicating possible problems in the integration of new cells at the interface between the old, intact tissue and the regenerated portion of the spinal cord. Taken together, the results of the present study are consistent with the hypothesis that apoptosis plays a role in the development of the new tissue during the cell replacement phase of the regenerating teleostean spinal cord. © 2009 Elsevier B.V. All rights reserved.
1.
Introduction
Teleost fish exhibit a remarkable degree of structural and functional regeneration after central nervous system (CNS) injury during adulthood (Becker and Becker, 2008; Zupanc, 2008a,b, 2009). This contrasts with mammals in which CNS lesions lead to severe and irreparable damage (Profyris et al.,
2004; Johansson, 2007; Bramlett and Dietrich, 2007). In mammalian systems, CNS injury causes high levels of necrosis within the first few days, leading to inflammation and, ultimately, to cavitation (Beattie et al., 2002; Profyris et al., 2004; Fitch and Silver, 2008). By contrast, in the CNS of teleost fish apoptosis is the predominant type of cell death immediately after injury (Zupanc et al., 1998; Sîrbulescu et al., 2009),
⁎ Corresponding author. School of Engineering and Science, Jacobs University Bremen, P.O. BOX 750 561, 28725 Bremen, Germany. Fax: +49 421 200 49 3244. E-mail address: [email protected] (G.K.H. Zupanc). Abbreviations: BrdU, 5-bromo-2′-deoxyuridine; BSA, bovine serum albumin; CNS, central nervous system; GFAP, glial fibrillary acidic protein; PBS, phosphate-buffered saline; RT, room temperature; SEM, standard error of the mean; TBS, tris-buffered saline 0006-8993/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.brainres.2009.09.071
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thus likely reducing tissue inflammation. In the weakly electric fish Apteronotus leptorhynchus, one of the most intensively studied teleostean model systems, apoptotic cell death peaks several hours after injury and rapidly declines 2– 5 days later in both the brain and the spinal cord (Zupanc et al., 1998; Sîrbulescu et al., 2009). This first stage of wound repair, characterized by the elimination of injured cells (arbitrarily termed “cell elimination phase”), is followed by the development of new tissue (arbitrarily termed “cell replacement phase”). In the brain of A. leptorhynchus, the vast majority of new cells are generated 1– 10 days after a lesion, with the maximum proliferative activity occurring at 5 days (Zupanc and Ott, 1999). In the spinal cord of A. leptorhynchus, cell proliferation starts to increase 5 days after the amputation of the tail, and persists at high levels for at least 50 days (Sîrbulescu et al., 2009). Previous analysis has indicated that, in the spinal cord, apoptosis persists during the cell replacement phase at levels similar to baseline in intact tissue (Sîrbulescu et al., 2009). However, virtually nothing is known about the spatiotemporal pattern of occurrence of apoptotic cell death within the regenerating nervous tissue. In the present study, we have, therefore, examined apoptotic cell death during the cell replacement phase up to 210 days after caudal spinal cord transection in A. leptorhynchus. Using immunolabeling against active caspase-3, an executioner caspase activated in late stages of apoptosis, we first examined the occurrence of apoptosis among newly generated cells, as identified by incorporation of the thymidine analogue 5-bromo-2′-deoxyuridine (BrdU). In a second set of experiments, we characterized the spatio-temporal dynamics of apoptotic cell death in the regenerating spinal cord. In the third and final part, we determined the identity of the newly generated cells undergoing apoptosis by employing the neuronal marker Hu C/D, an RNA-binding protein expressed early during neuronal differentiation (Marusich and Weston, 1992), and the astrocytespecific marker glial fibrillary acidic protein (GFAP) in triple immunolabeling experiments. Taken together, these results have given indication of possible functions of apoptosis during spinal cord regeneration.
2.
Results
2.1.
Apoptosis of newly generated cells
In order to examine the relationship between apoptotic cells and new cells generated in response to injury, we analyzed the co-localization pattern of active caspase-3 and the mitotic marker BrdU. Spinal cord injury was induced by a 1-cm tail amputation. The thymidine analogue BrdU was injected intraperitoneally 10 days later, when maximum levels of cell proliferation occur (Sîrbulescu et al., 2009). The fish were perfused at post-injection intervals ranging between 2 h and 200 days, and the caudalmost 1 cm of the spinal cord was analyzed. At all time points, caspase-3+ cells were present in low numbers (mean: 1.2 cells per section ± SEM: 0.13 cells per section), corresponding to the baseline levels observed in intact spinal cord (Sîrbulescu et al., 2009). Double-labeled cells,
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positive for both active caspase-3 and BrdU, could be observed as early as 2 h after the BrdU injection (Fig. 1) and at all other post-BrdU-administration survival times examined. At 2 h post-injection, approximately 2% of the BrdU+ cells were also caspase-3+. This percentage increased to approximately 6% at 1–3 days, and reached a relative plateau at 17% (±SEM: 5%) between 5 and 100 days post-injection. At later time points, the proportion of double-labeled cells markedly decreased to 3% at 150 days and 1% at 200 days post-injection. Conversely, the proportion of caspase-3+ cells that were also BrdU+ was low at 2 h (19%), increased at later time points to reach up to 93% at 100 days post-injection, and then gradually decreased. These results indicate that the majority of apoptotic cells in the regenerating spinal cord were newly formed cells.
2.2. Radial distribution of caspase-3+/BrdU+ double-labeled cells To examine the radial distribution of caspase-3+/BrdU+ double-labeled cells within transverse spinal cord sections, we first drew a line through the center of the central canal and the center of any double-labeled cell, and extended this line up to the pial surface. We then measured both the distance from the central canal to the respective double-labeled cell along the line, and the distance between central canal and pial surface. The latter measurement was used to normalize for the variation in size or shape between different sections. A total of 1216 measurements were taken on sections from 12 individual fish at various time points during regeneration. To standardize the measured locations of double-labeled cells, we divided a typical spinal cord section into ten concentric regions of equal width (Fig. 2A) and determined the area of each of these regions. A histogram of the ratios between the experimentally determined number of cells in each region and the area of each region determined in a typical section showed that the areal density of cells was not uniform (Fig. 2B). Two-sample Kolomogorov–Smirnov tests between each pair of distributions indicated no significant differences between distributions at different time points. When all data were collapsed across time points, the distribution of normalized radii was significantly different from the one expected if the cells would have been uniformly distributed throughout a section (two-sample Kolomogorov–Smirnov test, Z = 4.968, p < 0.001). While a uniform distribution implies the same areal density of cells in all regions, measurements indicated that in the proximity of both the central canal and the pial surface of the spinal cord sections the numbers of caspase-3+/BrdU+ cells were significantly lower. In contrast, in mid-parenchymal regions (labeled 4–8) the areal density of double-labeled cells was higher than expected for a uniform distribution (Fig. 2B).
2.3. Longitudinal distribution of caspase-3+/BrdU+ double-labeled cells To qualitatively examine the longitudinal distribution of apoptotic cells among the new cells, we combined anticaspase-3 and anti-BrdU immunolabeling in sagittal spinal cord sections taken from fish at 50 days post-amputation and
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Fig. 1 – Double immunolabeling against BrdU and active caspase-3 in a transverse section of the regenerating spinal cord of A. leptorhynchus 10 days post-amputation and 2 h post-BrdU administration. In addition to double-labeled cells (A–C; filled arrowheads), several single-labeled caspase-3+ (arrows in A, C) and BrdU+ cells (open arrows in B, C) could be observed. Z-stack confocal imaging confirmed the presence of the immunolabel throughout these cells (C). Red and green lines in C indicate the level of the z-stack cross-section shown on the sides of the panel. The central canal is indicated by c. Bar = 100 μm.
Fig. 2 – Spatio-temporal distribution of double-labeled caspase-3+/BrdU+ cells within transverse sections of the regenerating spinal cord. The fish received an intraperitoneal BrdU injection 10 days after tail amputation and were killed at various post-BrdU administration survival times. (A) Typical spinal cord section divided into 10 concentric regions of equal width. Bar = 50 μm. (B) Probability of encountering a double-labeled cell within a given region between the central canal and the pial surface of a spinal cord section at progressive post-BrdU administration survival times. In the proximity of the central canal and close to the pial surface, the areal density of double-labeled cells was lower than expected for a uniform distribution (dotted line). In contrast, in mid-parenchymal regions this probability was higher than expected, assuming a uniform distribution. No significant differences were observed between the distributions at different post-BrdU administration survival times.
40 days post-BrdU injection (Figs. 3A–C). At this survival time, approximately 3 mm of spinal cord had regenerated, a value in agreement with previous estimates (Sîrbulescu et al., 2009). In the section shown, a region of BrdU+ proliferating cells was evident caudal to the site of the lesion, between 1700 and 2300 μm rostrally from the caudal tip of the spinal cord (Figs. 3D–F), whereas an area with elevated numbers of apoptotic cells was located between 3400 and 3900 μm from the caudal
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Fig. 3 – Spatial distribution of caspase-3+, BrdU+, and double-labeled cells within longitudinal sections of the regenerated spinal cord 50 days post-amputation and 40 days post-BrdU administration. (A–C) At this time point, approximately 3 mm of the spinal cord had regenerated. Part of the regenerated portion of the spinal cord, and a region adjacent to the lesion site, indicated by dotted lines, are shown at higher magnification in panels D–F and G–I, respectively. (D–F) Within the regenerated portion of the spinal cord, numerous BrdU+ cells (open arrows in E, F), but relatively few caspase-3+ cells (short arrows in D, F), are found. Double-labeled cells are extremely rare in the regenerate, and none is present in the image shown. In the spinal cord tissue adjacent to the lesion site, increased numbers of caspase-3+ cells were present (G). Moreover, many of these apoptotic cells were also BrdU+ (H, I; arrowheads). This indicates that it is new cells that preferentially undergo apoptosis, and that the vast majority of these apoptotic events take place at the interface between regenerated and intact tissue. Caudal is towards left, dorsal upwards. Bar = 200 μm in A–C and 100 μm in D–I.
tip of the spinal cord, immediately rostral to the initial lesion site. Notably, in the newly developed tissue there was little double-labeling, except in areas close to the lesion site. In the latter areas, which were distinguished by their abundance of
apoptotic cells, the majority of caspase-3+ cells were also BrdU+ (Figs. 3G–I). For a quantitative determination of the spatiotemporal dynamics of cell division and apoptosis along the longitudinal
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Fig. 4 – Longitudinal distribution of BrdU+, caspase-3+ and double-labeled cells within the caudalmost 1 cm of the regenerating spinal cord. (A–K) A single BrdU injection was administered 10 days after spinal cord amputation, and the fish were perfused after various survival times, as indicated. A pronounced peak in the number of proliferating cells (grey line) was evident in the proximity of the initial lesion site (indicated by arrows) at all time points. The apparent shift in the location of the BrdU+ peak with increasing post-BrdU administration survival times reflects subsequent growth of the spinal cord. Caspase-3+ cell levels (dotted line), although much lower overall, showed a similar peak located approximately 500 μm rostrally to the cell proliferation peak, up to 100 days post-injection (A–I). At later time points, the number of caspase-3+ cells was very low, and their distribution appeared to be uniform throughout the 1-cm tissue block analyzed (J, K). A large proportion of the caspase-3+ cells were also BrdU+ (black line). Each of the graphs is a two-point average plot of one individual for each survival time.
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axis, we analyzed consecutive transverse sections along a 1cm segment of the regenerating spinal cord at post-injection intervals ranging between 2 h and 200 days (n = 22 fish). The analyzed region always included the complete regenerate, plus an intact portion of the spinal cord and the interface between the two. In all analyzed fish from 2 h up to 100 days post-injection survival (n = 18), a peak in the number of BrdU+ cells and another peak in the number of active caspase-3+ cells could be observed along the longitudinal axis (Figs. 4A–I). At 150 and 200 days post-injection, the peak of apoptotic cells disappeared, whereas the peak of BrdU+ cells persisted (Figs. 4J, K). Although the absolute magnitude of these peaks varied between time points and individuals, a similar pattern was evident in all cases. At short post-injection survival times (up to 15 days post-amputation/5 days post-BrdU injection), the high density of BrdU+ cells started at, or immediately next to, the lesion site, extending up to approximately 1000 μm rostrally. A peak of apoptotic cells, usually much smaller in magnitude, followed at approximately 500–1500 μm rostral to the lesion site. Numerous apoptotic cells were double-labeled. Interestingly, at later time points, the cell proliferation peak could be observed increasingly farther from the caudal tip of the regenerating spinal cord, up to 4000–6000 μm at 100– 200 days post-injection (Figs. 4I–K). Since the spinal cord continues to grow caudally, the fact that the BrdU peak remained relatively stationary indicated that few of the dividing cells labeled at the time of the BrdU injection migrated caudally over long distances. The apoptosis peak was always seen rostral to the proliferation peak, which was very close to the initial injury site. Fig. 5 schematically shows the regrowth of the spinal cord and the distribution of caspase-3+, BrdU+, and double-labeled cells relative to the site of the lesion.
2.4. cells
Apoptosis after differentiation of newly generated
To examine the relationship between apoptotic cells and differentiated cells among the newly generated ones, we combined immunolabeling against active caspase-3 and BrdU with labeling against the early neuronal marker Hu (Marusich and Weston, 1992) or the astroglial marker GFAP (Eng et al., 2000). Examples of such labeling are shown in Figs. 6 and 7.
2.4.1.
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Fig. 5 – Tail amputation paradigm and spinal cord regeneration after the lesion. (A) A. leptorhynchus with the location of the spinal cord (dotted line). The site where the tail was amputated is indicated by the dashed line. Ten days after the amputation, a single dose of BrdU was administered through intraperitoneal injection, and the fish were killed at various post-BrdU-administration survival times. (B) Schematic representation of the regenerating spinal cord. BrdU application labels numerous proliferating cells in the vicinity of the lesion at all post-amputation survival times (red dots). The majority of BrdU+ cells remain localized at the site of origin, but a few migrate caudally into newly generated tissue. Apoptotic caspase-3+ cells (green dots) are located mostly at the site of the injury, where they persist at levels above-baseline up to at least 110 days after the injury. At the lesion site, most of the caspase-3+ cells are also BrdU+ (black dots), indicating that most of the cells that undergo apoptosis at this location are newly generated ones. Baseline levels of caspase-3+ and BrdU+ cells can be observed in the intact portion of the spinal cord (shown to the right of the transection site). The dashed line indicates the site of transection. Bars = 1 cm in A, 1 mm in B.
Active-caspase-3/BrdU/Hu triple labeling
Immunolabeling confirmed the presence of large numbers of newly differentiated BrdU+/Hu+ neurons in the regenerating A. leptorhynchus spinal cord. Some of these new neuronal cells underwent apoptosis, as indicated by triple caspase-3+/ BrdU+/Hu+ immunostaining (Fig. 6). To quantitatively analyze the occurrence of apoptosis among the cell types, we determined the total number of caspase-3+ cells in spinal cord sections from fish treated as described above, while checking each such cell for double- or triple-labeling against BrdU, Hu, or both (Fig. 8A). This analysis indicated that neurons died at relatively constant rates (0.5 ± SEM: 0.04 cells per section) up to 100 days post-injection, comprising approximately 38% (±SEM: 3%) of apoptotic cells. At 150 days, this value decreased to 0.09 cells
per section (25% of apoptotic cells), while at 200 days only 0.006 cells per section (2% of dying cells) expressed neuronal markers. Triple-labeled caspase-3+/BrdU+/Hu+ cells were observed, to various degrees, at all survival times except at 200 days (Fig. 8A). While at 2 h post-injection only 0.14 cells per section were triple-labeled (8% of the total number of apoptotic cells), this value reached 0.3 cells per section (16% of apoptotic cells) at 1–3 days, and remained relatively constant at 0.3 ±SEM: 0.03 cells per section (25% ±SEM: 2% of apoptotic cells) between 5 and 100 days post-injection. At 150 days post-injection triple-labeled cells amounted to only 0.04 cells per section (10% of apoptotic cells), while at 200 days post-injection no triple-labeled cells could be observed. It
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Fig. 6 – Apoptotic cell death of newly differentiated neurons in the regenerated spinal cord 40 days post-amputation and 30 days post-BrdU administration. (A–D) Triple immunolabeling against active caspase-3, BrdU, and the neuronal marker Hu C/D revealed that some of the newly generated neurons (BrdU+/Hu+; short arrows) are also caspase-3+ (filled arrowheads). Such triple-labeled cells were often found at peripheral locations within spinal cord sections. Apoptotic neurons that were not BrdU+ were also observed (open arrowheads). Red and green lines in D indicate the level of the z-stack cross-section shown on the sides of the panel. The section was taken from the regenerated part of the spinal cord. The central canal is indicated by c. Bar = 100 μm.
should be noted that, due to the administration of a single dose of BrdU after amputation, it is evident that the absolute numbers of new neuronal cells and, consequently, of apoptotic new neurons, are underestimated in the results shown here.
2.4.2.
Active-caspase-3/BrdU/GFAP triple labeling
Similarly to neurons, a certain portion of the newly generated BrdU+/GFAP+ astrocytes died via apoptosis in the regenerating spinal cord (Fig. 7). When analyzed quantitatively, caspase-3+/ GFAP+ astroglia levels remained relatively constant at 0.3 ± SEM: 0.05 cells per section, (24% ±SEM: 1% of apoptotic cells) at any of the analyzed time points, indicating the existence of a definite baseline level of astrocytic apoptosis in the regenerating spinal cord (Fig. 8B). Triple-labeled caspase-3+/BrdU+/GFAP+ cells were observed in very low numbers as early as 2 h after a BrdU injection, when they reached only 0.03 cells per section (1% of all apoptotic cells). The numbers of triple-labeled cells
increased to 0.18 ± SEM: 0.015 cells per section (14% ±SEM: 1.5% of apoptotic cells) between 1 and 20 days post-injection, and then decreased slightly to approximately 0.12 ± SEM: 0.04 cells per section (9% ± SEM: 2.6% of apoptotic cells) between 30 and 100 days. At 150 days, only 0.01 cells per section were triple-labeled (3% of the apoptotic cells), while at 200 days no such cells could be found anymore, further confirming a return to baseline levels in terms of differentiation and integration of new cells (Fig. 8B).
3.
Discussion
3.1. Apoptosis: regulatory function during spinal cord regeneration? Previous work has demonstrated the enormous potential of A. leptorhynchus as a model of successful regeneration of the adult CNS after injury (for reviews, see Zupanc 2008a,b, 2009).
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Fig. 7 – Triple immunolabeling against caspase-3, BrdU, and the astroglial marker GFAP 40 days post-amputation and 30 days post-BrdU administration. (A–D) New astrocytes (BrdU+/GFAP+) were observed throughout spinal cord sections (arrows). Some of these new cells were also caspase-3+ (filled arrowheads). Apoptotic astrocytes that were not BrdU+ were also observed (A, C, D, open arrowheads). Red and green lines in D indicate the level of the z-stack cross-section shown on the sides of the panel. The section was taken from the regenerated part of the spinal cord. The central canal is indicated by c. Bar = 100 μm.
This ability has been documented in both the brain and the spinal cord. For example, a stab wound lesion in the cerebellum of this species has been shown to heal within a few weeks (Zupanc et al., 1998; Zupanc and Ott, 1999). Similarly, within 5 weeks after amputation of the caudal portion of the spinal cord the fish show complete structural and functional regeneration (Sîrbulescu et al., 2009). This high regenerative potential is made possible by massive cell proliferation induced by the injury, and by successful differentiation and integration of the new cells into the existing neural network (Zupanc and Ott, 1999; Sîrbulescu et al., 2009). A second factor that may facilitate wound healing is apoptosis. This type of cell death is likely to minimize tissue inflammation during elimination of damaged cells at the lesion site (Zupanc et al., 1998; Sîrbulescu et al., 2009). Furthermore, the results of the present study are also consistent with the notion that apoptotic cell death might be involved in the development of new tissue. A functional role of apoptosis during the formation of new tissue is supported by observations in a variety of organisms,
ranging from invertebrates to mammals. For example, apoptosis has been shown to act as a mechanism for regulating neuronal connectivity (for reviews, see Chan et al., 2002; Buss et al., 2006). Moreover, in adult organisms in which cell proliferation persists either throughout the CNS or in restricted areas, apoptosis appears to ensure that supernumerary cells are eliminated (Buss et al., 2006). After amputation of the caudal spinal cord of A. leptorhynchus, complete re-growth of nervous tissue occurs, as it does during normal development. It is, therefore, likely that apoptosis after the initial cell elimination phase also plays a role in the structuring of new tissue and in ensuring proper integration of the new cells. In support of this notion, we have found in the present study that up to 110 days post-amputation approximately 38% of the observed apoptotic cells were differentiated neurons, while approximately 24% were GFAP+ astrocytes. These results indicate that a large portion of cells die during later stages of development, presumably at their target areas. This is in agreement with earlier findings in the cerebellum of A. leptorhynchus, where approximately 50% of the newly
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Fig. 8 – Quantitative analysis of caspase-3+ cells in the caudalmost 1 cm of spinal cord after triple-immunolabeling against caspase-3, BrdU, and Hu C/D (A) or GFAP (B). BrdU was administered intraperitoneally 10 days after tail amputation. (A) Analysis indicated that up to 100 days post-BrdU administration survival, neurons died at a relatively constant rate, comprising on average 38% (±SEM: 3%) of apoptotic cells. Newly differentiated neurons (caspase-3+/BrdU+/Hu+) constituted 8% of apoptotic cells at 2 h post-BrdU administration, increased to 16% at 1–3 days, and then maintained a relatively constant level of approximately 25% (±SEM: 2%) of apoptotic cells between 5 and 100 days post-BrdU administration. After 100 days, these percentages decreased, with neurons representing only 2% of apoptotic cells. No apoptotic new neurons were observed at 200 days. (B) Astrocytes constituted on average 24% (±SEM: 1%) of apoptotic cells at all time points analyzed. Apoptotic newly differentiated astroglia (caspase-3+/BrdU+/GFAP+) were present at very low levels (1%) at 2 h post-injection. Their numbers increased to 14% (±SEM: 1.5%) between 1 and 20 days, followed by a decline to approximately 9% (±SEM: 2.6%) between 30 and 100 days, and to 3% by 150 days post-injection. At 200 days post-injection no triple-labeled cells could be observed.
generated cells die upon arrival at the target sites, presumably by undergoing apoptosis (Soutschek and Zupanc, 1996; Ott et al., 1997). Is there a selection process that determines which cells undergo apoptosis? A previous study in the intact brain of A.
leptorhynchus has shown that apoptosis affects preferentially those adult-born cells that exhibit numerical chromosome aberrations due to mitotic segregation defects (Rajendran et al., 2008). It is likely that aneuploidy also occurs among the new cells generated in response to injury, because they derive from the same pool of stem cells as the new cells in the intact brain (Zupanc and Ott, 1999). It would, therefore, be interesting to examine whether part of the new cells generated after spinal cord injury are also aneuploid, and if so, whether these cells are eliminated with higher frequency through apoptosis than euploid cells. Interestingly, we observed an uneven radial distribution of newly generated cells undergoing apoptosis within the regenerating spinal cord. Such cells were present in relatively low numbers in the proximity of the central canal and the pial surface (Fig. 2B), suggesting a protective effect in these areas. In other regeneration-competent organisms, such as urodele amphibians, ependymal cells have been shown to upregulate trophic factors, such as retinoic acid and fibroblast growth factor, after spinal cord lesion (Chernoff et al., 2003; Ferretti et al., 2003). Furthermore, fibroblast growth factor has been demonstrated to effectively inhibit caspase-3 activation in vitro (Miho et al., 1999). Since an ependymal response to injury also occurs in A. leptorhynchus, it is possible that local diffusion of growth factors might inhibit apoptosis in the proximity of the central canal. Why apoptosis levels are relatively low close to the pial surface remains unclear, but perhaps the glia limitans exerts a similar trophic function as do the ependymal cells at the central canal. Another line of evidence in favor of the restructuring function of apoptosis during the cell replacement phase of spinal cord regeneration is the observed caudo-rostral gradient of caspase-3+ cells. While very few caspase-3+ or doublelabeled caspase-3+/BrdU+ cells were found in the caudalmost (young) regions of the regenerated spinal cord, the number of such cells gradually increased, relative to baseline levels, in more rostral (older) parts (Fig. 4). Corroborated by the fact that spinal cord differentiation after tail amputation takes place in a rostro-caudal direction (Waxman and Anderson, 1986; Chernoff et al., 2003; Ferretti et al., 2003), this finding indicates that most apoptotic events do not occur at the site of cell proliferation, but instead in regions of cell maturation. In addition, while apoptotic cells constituted, on average, 15% of the BrdU+ cells in the period of 1–100 days after BrdU administration, between 60 and 90% of apoptotic cells were also BrdU+ during the same interval. This result demonstrates that the majority of apoptotic cells found throughout the analyzed portion of the spinal cord were, indeed, young cells generated after tail amputation. Taken together, these results support the hypothesis that apoptosis plays a regulatory role during nervous tissue regeneration in the spinal cord of A. leptorhynchus, presumably by eliminating superfluous or poorly integrated cells.
3.2. Elevated levels of apoptosis at the transition zone between intact and regenerated tissue After the initial phase of cell elimination by apoptosis, the number of apoptotic cells remained low throughout the intact, as well as the regenerated parts of the spinal cord, with one
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exception. At the site where the spinal cord had been cut, a pronounced peak of apoptotic cells, sometimes three times higher than baseline levels, could be observed up to 110 days after amputation (Fig. 4). Such long-term persistence of a relatively high frequency of occurrence of apoptotic cell death at this specific location may reflect a defective, or incomplete, organization of the transition zone between the intact tissue and the newly generated one. This finding is especially remarkable since the regenerated part of the spinal cord does receive innervation from intact regions (Sîrbulescu et al., 2009). It is possible that the damage inflicted by the lesion induces a long-lasting alteration of the local environment at the interface between the intact part and the regenerate, which significantly delays successful integration, but not differentiation, of new cells. The distribution of triple-labeled caspase-3+/BrdU+/Hu+ and caspase-3+/BrdU+/ GFAP+ cells at this location mirrored the distribution of caspase-3+ cells, indicating that the observed apoptotic peak included differentiated neurons and glia, and could be located at an integration interface within the regenerating spinal cord. Interestingly, a previous study in a related teleost species, which is capable of regenerating its tail after amputation similarly to A. leptorhynchus, has also revealed structural modifications at the transition zone between regenerated and unregenerated spinal cord (Anderson et al., 1984). In the latter study, longitudinal sections through spinal cord have shown a clear constriction of the tissue around the transition area, 4 months after amputation. Albeit not of the same magnitude, we have also observed some decrease in tissue density in transverse spinal cord sections at this level in A. leptorhynchus, suggesting that ongoing apoptosis exceeds cell proliferation in this area, and thus affects the structure of the tissue. In the present study, we monitored apoptotic cell death starting 10 days post-lesion. At this time point, the number of apoptotic cells at the site of injury was only one tenth of the levels found immediately after the amputation when massive elimination of cells through apoptosis occurs (Sîrbulescu et al., 2009). Furthermore, the peak of apoptosis regressed after 110 days post-amputation and was not visible anymore at 160 or 210 days after the lesion (Fig. 4). These results indicate that the long-lasting disruption of tissue at the original site of the cut is eventually resolved, and successful integration is finally achieved. At the same time, 150–200 days post-BrdU administration, the proportion of newly differentiated neurons and astroglia that undergo apoptosis becomes markedly lower (Fig. 8). This might represent the end of the integration process of the new cells, and the return of the tissue to a cytoarchitecture reminiscent of the pre-injury structure.
3.3.
New roles of apoptosis: a comparative view
In mammals, apoptotic cell death takes place not only during the acute phase associated with the primary injury, but also during secondary injury which extends for long periods of time after the initial insult (for reviews, see Beattie et al., 2002; Kwon et al., 2004; Profyris et al., 2004; Fitch and Silver, 2008; Johansson, 2007; Ling and Liu, 2007). Elevated levels of apoptosis during the secondary injury can last for many
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weeks after the lesion (Beattie et al., 2002; Kwon et al., 2004; Keane et al., 2006; Bramlett and Dietrich, 2007). Such high levels of cell death are considered one of the main reasons for the significant loss of neural tissue after spinal cord injury (Kim et al., 2003; Martiñón and Ibarra, 2008), thus seemingly supporting the notion of a detrimental role of apoptosis after injury in non-regenerating systems. By contrast, in regeneration-competent systems there is evidence of a rather supportive function of apoptosis after injury. During the first few days after experimentally applied lesions, apoptosis appears to mediate the “clean” elimination of damaged cells in both the brain (Zupanc et al., 1998) and the spinal cord of A. leptorhynchus (Sîrbulescu et al., 2009). As suggested by the present study, apoptosis might also be involved in tissue repair during the later stages of spinal cord regeneration by regulating the survival of newly formed cells. This is consistent with the observation in larval lampreys where apoptotic cell death persists for at least 12–16 weeks after spinal cord transection (Shifman et al., 2008). In Xenopus laevis tadpoles, inhibition of apoptosis has been shown to effectively prevent regeneration after tail transection (Tseng et al., 2007). Interestingly, in rats recent evidence indicates that preventing apoptosis for an extended duration after spinal cord injury may impair recovery and lead to an increase in necrosis and inflammation (Cittelly et al., 2008). These studies suggest that apoptosis does not only exert a negative function after a lesion by eliminating cells, as traditionally assumed. Instead, it may also play an important role for the development of new cells that replace tissue lost to injury. The spinal cord of A. leptorhynchus will provide an excellent regeneration-competent model system to test this hypothesis.
4.
Experimental procedures
4.1.
Animals
Fish were purchased from a tropical fish importer and kept in individual aquaria under defined conditions (Zupanc et al., 2006). A total of 25 adult fish with a total body length between 102 mm and 154 mm (mean: 126 mm; median: 126 mm) and body mass between 1.9 g and 8.2 g (mean: 4.1 g; median: 4.5 g) were used in this study. Sixteen fish were males and nine females, as revealed by post-mortem gonadal inspection. The gonadosomatic index, determined as fresh weight of gonads divided by body mass, ranged between 0.0013 and 0.005 (mean: 0.0032; median: 0.0030) in males, and between 0.0068 and 0.1841 (mean: 0.0904; median: 0.0940) in females. All experiments were carried out in accordance with the relevant German law, the Deutsches Tierschutzgesetz of 1998.
4.2.
Immunohistochemistry
Under general anesthesia with urethane (Sigma, Taufkirchen, Germany) and after application of 2% lidocaine (Euro OTC Pharma, Hannover, Germany) as a local anesthetic, 1 cm of the tail, measured rostrally from the white caudal ring, was amputated (n = 25 fish). After a 10-day recovery period, the
24
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fish received a single intraperitoneal injection of approximately 50 μl of a 3 mg/ml BrdU solution (Amersham/GE Healthcare, Freiburg, Germany). Following various post-amputation survival times (2 h, 1 day, 3 days, 5 days, 10 days, 20 days, 30 days, 40 days, 50 days, 100 days, 150 days, 200 days; n = 2 fish per survival time), animals were deeply anesthetized using ethyl 3-aminobenzoate methanesulfonate (MS-222; Sigma) and intracardially perfused. The 1-cm caudalmost segment was removed from the tail, post-fixed, and cryoprotected as described (Zupanc et al., 2005). Transverse or sagittal sections were cut at a thickness of 16 μm and mounted serially onto SuperFrost Plus Gold slides (Menzel-Gläser, Braunschweig, Germany).
4.2.1.
Active caspase-3/BrdU double immunolabeling
After the sections had been dried in a desiccator for 90 min at room temperature (RT), they were rinsed three times for 10 min each in 0.1 M tris-buffered saline (TBS) and blocked for 60 min with 3% normal sheep serum, 1% teleostean gelatin, 1% bovine serum albumin (BSA), and 0.3% Triton X-100 in TBS (all from Sigma). Sections were incubated with a purified monoclonal rabbit anti-active caspase-3 antibody (clone C92-605; Cat. No. 559565; BD Pharmigen, Heidelberg, Germany), diluted 1:100 in blocking solution, at 4 °C overnight. After three washes in TBS for 5 min each, the sections were blocked for 30 min at RT with 3% goat serum (Sigma), 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100, followed by incubation for 90 min at RT with the secondary antibody Alexa Fluor 488conjugated goat anti-rabbit IgG (Molecular Probes/Invitrogen, Karlsruhe, Germany), diluted 1:200 in blocking solution. After three washes in TBS for 10 min each, the sections were first incubated with a solution of 20% acetic acid and 70% ethanol in ddH2O for 5 min at RT, and then in 70% ethanol for 25 min at −20 °C. After three washes in TBS for 5 min each, the sections were incubated in 2 M HCl at 37 °C for 30 min, followed first by two rinses in 0.1 M borate buffer (pH 8.5) at RT for 20 min each, and then by six rinses in TBS for 5 min each. After blocking for 30 min with 3% normal sheep serum, 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100 in TBS, the sections were incubated with rat anti-BrdU antibody (Cat. No. OBT0030; Oxford Biotechnology through Biozol, Eching, Germany), diluted 1:200 in blocking solution, at 4 °C overnight. Following three rinses in TBS for 5 min each, the sections were blocked for 30 min at RT with 3% donkey serum (Sigma), 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100, and incubated for 90 min at RT with the secondary antibody, donkey anti-rat IgG conjugated to Cy3 (Jackson Immunoresearch through Dianova, Hamburg, Germany), at a dilution of 1:1500 in blocking solution. After three washes in TBS for 10 min each, the sections were embedded in polyvinyl alcohol containing n-propyl gallate.
4.2.2.
Active caspase-3/BrdU/Hu C/D triple immunolabeling
After drying in a desiccator for 90 min at RT, the sections were rinsed three times for 10 min each in 0.1 M phosphatebuffered saline (PBS), incubated for 30 min in 50 mM Tris, pH 8.0, and finally rinsed three times in PBS for 5 min each. Blocking was performed for 60 min with 3% normal sheep serum, 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100 in PBS, after which sections were incubated with a monoclonal
mouse anti-Hu C/D antibody (clone 16A11; Cat. No. A-21271; Molecular Probes), diluted 1:20 in blocking solution, at 4 °C overnight. After three washes in PBS for 5 min each, the sections were blocked for 30 min at RT with 3% goat serum, 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100 in PBS, followed by incubation for 90 min at RT with the secondary antibodies Alexa Fluor 635-conjugated goat anti-mouse IgG (Molecular Probes), diluted 1:200, and Coumarin AMCAconjugated goat anti-mouse IgG (Jackson Immunoresearch), diluted 1:100, in blocking solution. Further labeling for active caspase-3 and BrdU was performed as described above.
4.2.3.
Active caspase-3/BrdU/GFAP triple immunolabeling
After drying in a desiccator for 90 min at RT, the sections were rinsed three times for 10 min each in 0.1 M TBS, and blocked for 60 min with 3% normal sheep serum, 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100 in TBS. Sections were incubated with a monoclonal mouse anti-GFAP antibody (clone G-5-A; Cat. No. G3893; Sigma), diluted 1:50 in blocking solution, at 4 °C overnight. After three washes in TBS for 5 min each, the sections were blocked for 30 min at RT with 3% goat serum, 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100 in TBS, followed by incubation for 90 min at RT with the secondary antibodies Alexa Fluor 635conjugated goat anti-mouse IgG (Molecular Probes), diluted 1:200, and Coumarin AMCA-conjugated goat anti-mouse IgG (Jackson Immunoresearch), diluted 1:100, in blocking solution. Further labeling for active caspase-3 and BrdU was performed as described above.
4.3.
Antibody controls
Specificity of the rabbit anti-active caspase-3, rat-anti BrdU and mouse anti-GFAP primary antibodies in A. leptorhynchus was demonstrated previously both by pre-adsorption of the respective primary antibody with the specific antigen and by omission of the primary antibody during labeling (Clint and Zupanc, 2001; Sîrbulescu et al., 2009). The mouse anti-Hu C/D antibody used in the present study has previously been shown to recognize a single protein band at the expected molecular weight of 42 kDa in Western blots of tissue from teleostean CNS (Takeda et al., 2008).
4.4.
Microscopy and analysis
Sections were examined under a Zeiss Axioskop epifluorescence microscope (Carl Zeiss, Göttingen, Germany). Every complete section was analyzed, and all caspase-3-positive cells were counted. In double-labeling experiments, also all BrdU-positive cells were counted. In any experiment, a minimum of 64 sections were analyzed per fish. The location of labeled cells was entered into camera lucida drawings of spinal cord sections. Every caspase-3-positive cell was checked for double- or triple-labeling against other antigens. For confocal microscopy, specimens were examined on a Zeiss LSM 510 META laser-scanning microscope. Optical sections were taken with a pinhole opening of 1 Airy and at a resolution of 1024 × 1024 pixels, using LSM 5 (version 3.2; Carl Zeiss) software. For longitudinal sections, multiple consecutive images taken along the available portion of the spinal cord
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were digitally stitched using CorelDraw 11.0 (Corel Corporation, Ottawa, Canada) to give an overview of the reconstructed tissue section.
Acknowledgments This study received financial support from the Ernst A.-C. Lange-Stiftung and from Jacobs University Bremen.
REFERENCES
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Ling, X., Liu, D., 2007. Temporal and spatial profiles of cell loss after spinal cord injury: Reduction by a metalloporphyrin. J. Neurosci. Res. 85, 2175–2185. Martiñón, S., Ibarra, A., 2008. Pharmacological neuroprotective therapy for acute spinal cord injury: state of the art. Mini Rev. Med. Chem. 8, 222–230. Marusich, M.F., Weston, J.A., 1992. Identification of early neurogenic cells in the neural crest lineage. Dev. Biol. 149, 295–306. Miho, Y., Kouroku, Y., Fujita, E., Mukasa, T., Urase, K., Kasahara, T., Isoai, A., Momoi, M.Y., Momoi, T., 1999. bFGF inhibits the activation of caspase-3 and apoptosis of P19 embryonal carcinoma cells during neuronal differentiation. Cell Death Differ. 6, 463–470. Ott, R., Zupanc, G.K.H., Horschke, I., 1997. Long-term survival of postembryonically born cells in the cerebellum of gymnotiform fish, Apteronotus leptorhynchus. Neurosci. Lett. 221, 185–188. Profyris, C., Cheema, S.S., Zang, D., Azari, M.F., Boyle, K., Petratos, S., 2004. Degenerative and regenerative mechanisms governing spinal cord injury. Neurobiol. Dis. 15, 415–436. Rajendran, R.S., Wellbrock, U.M., Zupanc, G.K.H., 2008. Apoptotic cell death, long-term persistence, and neuronal differentiation of aneuploid cells generated in the adult brain of teleost fish. Dev. Neurobiol. 68, 1257–1268. Shifman, M.I., Zhang, G., Selzer, M.E., 2008. Delayed death of identified reticulospinal neurons after spinal cord injury in lampreys. J. Comp. Neurol. 510, 269–282. Sîrbulescu, R.F., Ilieş, I., Zupanc, G.K.H., 2009. Structural and functional regeneration after spinal cord injury in the weakly electric teleost fish, Apteronotus leptorhynchus. J. Comp. Physiol. A 195, 699–714. Soutschek, J., Zupanc, G.K.H., 1996. Apoptosis in the cerebellum of adult teleost fish, Apteronotus leptorhynchus. Dev. Brain Res. 97, 279–286. Takeda, A., Nakano, M., Goris, R.C., Funakoshi, K., 2008. Adult neurogenesis with 5-HT expression in lesioned goldfish spinal cord. Neuroscience 151, 1132–1141. Tseng, A.S., Adams, D.S., Qiu, D., Koustubhan, P., Levin, M., 2007. Apoptosis is required during early stages of tail regeneration in Xenopus laevis. Dev. Biol. 301, 62–69. Waxman, S.G., Anderson, M.J., 1986. Regeneration of central nervous system structures: Apteronotus spinal cord as a model system. In: Bullock, T.H., Heiligenberg, W. (Eds.), Electroreception. Wiley & Sons, New York, pp. 183–208. Zupanc, G.K.H., 2008a. Adult neurogenesis and neuronal regeneration in the brain of teleost fish. J. Physiol. Paris 102, 357–373. Zupanc, G.K.H., 2008b. Adult neurogenesis in teleost fish. In: Gage, F.H., Kempermann, G., Song, H.J. (Eds.), Adult Neurogenesis. Cold Spring Harbor Laboratory Press, Cold Spring Harbor/New York, pp. 571–592. Zupanc, G.K.H., 2009. Towards brain repair: insights from teleost fish. Semin. Cell Dev. Biol. 20, 683–690. Zupanc, G.K.H., Ott, R., 1999. Cell proliferation after lesions in the cerebellum of adult teleost fish: time course, origin, and type of new cells produced. Exp. Neurol. 160, 78–87. Zupanc, G.K.H., Kompass, K.S., Horschke, I., Ott, R., Schwarz, H., 1998. Apoptosis after injuries in the cerebellum of adult teleost fish. Exp. Neurol. 152, 221–230. Zupanc, G.K.H., Hinsch, K., Gage, F.H., 2005. Proliferation, migration, neuronal differentiation, and long-term survival of new cells in the adult zebrafish brain. J. Comp. Neurol. 488, 290–319. Zupanc, G.K.H., Sîrbulescu, R.F., Nichols, A., Ilies, I., 2006. Electric interactions through chirping behavior in the weakly electric fish, Apteronotus leptorhynchus. J. Comp. Physiol. A 192, 159–173.
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www.elsevier.com/locate/brainres
Research Report
Dynamics of caspase-3-mediated apoptosis during spinal cord regeneration in the teleost fish, Apteronotus leptorhynchus Ruxandra F. Sîrbulescu a , Günther K.H. Zupanc a,b,⁎ a
School of Engineering and Science, Jacobs University Bremen, P.O. BOX 750 561, 28725 Bremen, Germany Department of Biology, Northeastern University, 360 Huntington Avenue, Boston, MA 02115, USA
b
A R T I C LE I N FO
AB S T R A C T
Article history:
In contrast to mammals, adult teleost fish exhibit a vast potential for central nervous
Accepted 17 September 2009
system regeneration after injury. Among other mechanisms, this capacity is mediated by
Available online 24 September 2009
replacement of cells lost to injury by new neurons and glia. Here, we examined the spatio-
Keywords:
differentiation of new cells, during this cell replacement phase. As an experimental
temporal dynamics of apoptosis, and its relationship to the generation and the Spinal cord regeneration
paradigm, caudal transection of the spinal cord in the teleost fish Apteronotus leptorhynchus
Apoptosis
was used. During the cell replacement phase, there was a rather constant percentage of new
Cell proliferation
cells (identified by incorporation of 5-bromo-2′-deoxyuridine into newly synthesized DNA)
Adult neurogenesis
that underwent apoptosis (identified by anti-active caspase-3 immunolabeling). Many of
Caspase-3
these cells were also immunopositive for the marker proteins Hu C/D or glial fibrillary acidic protein, indicating that a large portion of cells undergo apoptosis after differentiation into neurons or glia, respectively. The spatial distribution of apoptotic cells was uneven, displaying a radial peak in the mid parenchymal regions and a longitudinal peak at the site of the initial spinal transection. The latter persisted for over 100 days post-injury, indicating possible problems in the integration of new cells at the interface between the old, intact tissue and the regenerated portion of the spinal cord. Taken together, the results of the present study are consistent with the hypothesis that apoptosis plays a role in the development of the new tissue during the cell replacement phase of the regenerating teleostean spinal cord. © 2009 Elsevier B.V. All rights reserved.
1.
Introduction
Teleost fish exhibit a remarkable degree of structural and functional regeneration after central nervous system (CNS) injury during adulthood (Becker and Becker, 2008; Zupanc, 2008a,b, 2009). This contrasts with mammals in which CNS lesions lead to severe and irreparable damage (Profyris et al.,
2004; Johansson, 2007; Bramlett and Dietrich, 2007). In mammalian systems, CNS injury causes high levels of necrosis within the first few days, leading to inflammation and, ultimately, to cavitation (Beattie et al., 2002; Profyris et al., 2004; Fitch and Silver, 2008). By contrast, in the CNS of teleost fish apoptosis is the predominant type of cell death immediately after injury (Zupanc et al., 1998; Sîrbulescu et al., 2009),
⁎ Corresponding author. School of Engineering and Science, Jacobs University Bremen, P.O. BOX 750 561, 28725 Bremen, Germany. Fax: +49 421 200 49 3244. E-mail address: [email protected] (G.K.H. Zupanc). Abbreviations: BrdU, 5-bromo-2′-deoxyuridine; BSA, bovine serum albumin; CNS, central nervous system; GFAP, glial fibrillary acidic protein; PBS, phosphate-buffered saline; RT, room temperature; SEM, standard error of the mean; TBS, tris-buffered saline 0006-8993/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.brainres.2009.09.071
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thus likely reducing tissue inflammation. In the weakly electric fish Apteronotus leptorhynchus, one of the most intensively studied teleostean model systems, apoptotic cell death peaks several hours after injury and rapidly declines 2– 5 days later in both the brain and the spinal cord (Zupanc et al., 1998; Sîrbulescu et al., 2009). This first stage of wound repair, characterized by the elimination of injured cells (arbitrarily termed “cell elimination phase”), is followed by the development of new tissue (arbitrarily termed “cell replacement phase”). In the brain of A. leptorhynchus, the vast majority of new cells are generated 1– 10 days after a lesion, with the maximum proliferative activity occurring at 5 days (Zupanc and Ott, 1999). In the spinal cord of A. leptorhynchus, cell proliferation starts to increase 5 days after the amputation of the tail, and persists at high levels for at least 50 days (Sîrbulescu et al., 2009). Previous analysis has indicated that, in the spinal cord, apoptosis persists during the cell replacement phase at levels similar to baseline in intact tissue (Sîrbulescu et al., 2009). However, virtually nothing is known about the spatiotemporal pattern of occurrence of apoptotic cell death within the regenerating nervous tissue. In the present study, we have, therefore, examined apoptotic cell death during the cell replacement phase up to 210 days after caudal spinal cord transection in A. leptorhynchus. Using immunolabeling against active caspase-3, an executioner caspase activated in late stages of apoptosis, we first examined the occurrence of apoptosis among newly generated cells, as identified by incorporation of the thymidine analogue 5-bromo-2′-deoxyuridine (BrdU). In a second set of experiments, we characterized the spatio-temporal dynamics of apoptotic cell death in the regenerating spinal cord. In the third and final part, we determined the identity of the newly generated cells undergoing apoptosis by employing the neuronal marker Hu C/D, an RNA-binding protein expressed early during neuronal differentiation (Marusich and Weston, 1992), and the astrocytespecific marker glial fibrillary acidic protein (GFAP) in triple immunolabeling experiments. Taken together, these results have given indication of possible functions of apoptosis during spinal cord regeneration.
2.
Results
2.1.
Apoptosis of newly generated cells
In order to examine the relationship between apoptotic cells and new cells generated in response to injury, we analyzed the co-localization pattern of active caspase-3 and the mitotic marker BrdU. Spinal cord injury was induced by a 1-cm tail amputation. The thymidine analogue BrdU was injected intraperitoneally 10 days later, when maximum levels of cell proliferation occur (Sîrbulescu et al., 2009). The fish were perfused at post-injection intervals ranging between 2 h and 200 days, and the caudalmost 1 cm of the spinal cord was analyzed. At all time points, caspase-3+ cells were present in low numbers (mean: 1.2 cells per section ± SEM: 0.13 cells per section), corresponding to the baseline levels observed in intact spinal cord (Sîrbulescu et al., 2009). Double-labeled cells,
15
positive for both active caspase-3 and BrdU, could be observed as early as 2 h after the BrdU injection (Fig. 1) and at all other post-BrdU-administration survival times examined. At 2 h post-injection, approximately 2% of the BrdU+ cells were also caspase-3+. This percentage increased to approximately 6% at 1–3 days, and reached a relative plateau at 17% (±SEM: 5%) between 5 and 100 days post-injection. At later time points, the proportion of double-labeled cells markedly decreased to 3% at 150 days and 1% at 200 days post-injection. Conversely, the proportion of caspase-3+ cells that were also BrdU+ was low at 2 h (19%), increased at later time points to reach up to 93% at 100 days post-injection, and then gradually decreased. These results indicate that the majority of apoptotic cells in the regenerating spinal cord were newly formed cells.
2.2. Radial distribution of caspase-3+/BrdU+ double-labeled cells To examine the radial distribution of caspase-3+/BrdU+ double-labeled cells within transverse spinal cord sections, we first drew a line through the center of the central canal and the center of any double-labeled cell, and extended this line up to the pial surface. We then measured both the distance from the central canal to the respective double-labeled cell along the line, and the distance between central canal and pial surface. The latter measurement was used to normalize for the variation in size or shape between different sections. A total of 1216 measurements were taken on sections from 12 individual fish at various time points during regeneration. To standardize the measured locations of double-labeled cells, we divided a typical spinal cord section into ten concentric regions of equal width (Fig. 2A) and determined the area of each of these regions. A histogram of the ratios between the experimentally determined number of cells in each region and the area of each region determined in a typical section showed that the areal density of cells was not uniform (Fig. 2B). Two-sample Kolomogorov–Smirnov tests between each pair of distributions indicated no significant differences between distributions at different time points. When all data were collapsed across time points, the distribution of normalized radii was significantly different from the one expected if the cells would have been uniformly distributed throughout a section (two-sample Kolomogorov–Smirnov test, Z = 4.968, p < 0.001). While a uniform distribution implies the same areal density of cells in all regions, measurements indicated that in the proximity of both the central canal and the pial surface of the spinal cord sections the numbers of caspase-3+/BrdU+ cells were significantly lower. In contrast, in mid-parenchymal regions (labeled 4–8) the areal density of double-labeled cells was higher than expected for a uniform distribution (Fig. 2B).
2.3. Longitudinal distribution of caspase-3+/BrdU+ double-labeled cells To qualitatively examine the longitudinal distribution of apoptotic cells among the new cells, we combined anticaspase-3 and anti-BrdU immunolabeling in sagittal spinal cord sections taken from fish at 50 days post-amputation and
16
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Fig. 1 – Double immunolabeling against BrdU and active caspase-3 in a transverse section of the regenerating spinal cord of A. leptorhynchus 10 days post-amputation and 2 h post-BrdU administration. In addition to double-labeled cells (A–C; filled arrowheads), several single-labeled caspase-3+ (arrows in A, C) and BrdU+ cells (open arrows in B, C) could be observed. Z-stack confocal imaging confirmed the presence of the immunolabel throughout these cells (C). Red and green lines in C indicate the level of the z-stack cross-section shown on the sides of the panel. The central canal is indicated by c. Bar = 100 μm.
Fig. 2 – Spatio-temporal distribution of double-labeled caspase-3+/BrdU+ cells within transverse sections of the regenerating spinal cord. The fish received an intraperitoneal BrdU injection 10 days after tail amputation and were killed at various post-BrdU administration survival times. (A) Typical spinal cord section divided into 10 concentric regions of equal width. Bar = 50 μm. (B) Probability of encountering a double-labeled cell within a given region between the central canal and the pial surface of a spinal cord section at progressive post-BrdU administration survival times. In the proximity of the central canal and close to the pial surface, the areal density of double-labeled cells was lower than expected for a uniform distribution (dotted line). In contrast, in mid-parenchymal regions this probability was higher than expected, assuming a uniform distribution. No significant differences were observed between the distributions at different post-BrdU administration survival times.
40 days post-BrdU injection (Figs. 3A–C). At this survival time, approximately 3 mm of spinal cord had regenerated, a value in agreement with previous estimates (Sîrbulescu et al., 2009). In the section shown, a region of BrdU+ proliferating cells was evident caudal to the site of the lesion, between 1700 and 2300 μm rostrally from the caudal tip of the spinal cord (Figs. 3D–F), whereas an area with elevated numbers of apoptotic cells was located between 3400 and 3900 μm from the caudal
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17
Fig. 3 – Spatial distribution of caspase-3+, BrdU+, and double-labeled cells within longitudinal sections of the regenerated spinal cord 50 days post-amputation and 40 days post-BrdU administration. (A–C) At this time point, approximately 3 mm of the spinal cord had regenerated. Part of the regenerated portion of the spinal cord, and a region adjacent to the lesion site, indicated by dotted lines, are shown at higher magnification in panels D–F and G–I, respectively. (D–F) Within the regenerated portion of the spinal cord, numerous BrdU+ cells (open arrows in E, F), but relatively few caspase-3+ cells (short arrows in D, F), are found. Double-labeled cells are extremely rare in the regenerate, and none is present in the image shown. In the spinal cord tissue adjacent to the lesion site, increased numbers of caspase-3+ cells were present (G). Moreover, many of these apoptotic cells were also BrdU+ (H, I; arrowheads). This indicates that it is new cells that preferentially undergo apoptosis, and that the vast majority of these apoptotic events take place at the interface between regenerated and intact tissue. Caudal is towards left, dorsal upwards. Bar = 200 μm in A–C and 100 μm in D–I.
tip of the spinal cord, immediately rostral to the initial lesion site. Notably, in the newly developed tissue there was little double-labeling, except in areas close to the lesion site. In the latter areas, which were distinguished by their abundance of
apoptotic cells, the majority of caspase-3+ cells were also BrdU+ (Figs. 3G–I). For a quantitative determination of the spatiotemporal dynamics of cell division and apoptosis along the longitudinal
18
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Fig. 4 – Longitudinal distribution of BrdU+, caspase-3+ and double-labeled cells within the caudalmost 1 cm of the regenerating spinal cord. (A–K) A single BrdU injection was administered 10 days after spinal cord amputation, and the fish were perfused after various survival times, as indicated. A pronounced peak in the number of proliferating cells (grey line) was evident in the proximity of the initial lesion site (indicated by arrows) at all time points. The apparent shift in the location of the BrdU+ peak with increasing post-BrdU administration survival times reflects subsequent growth of the spinal cord. Caspase-3+ cell levels (dotted line), although much lower overall, showed a similar peak located approximately 500 μm rostrally to the cell proliferation peak, up to 100 days post-injection (A–I). At later time points, the number of caspase-3+ cells was very low, and their distribution appeared to be uniform throughout the 1-cm tissue block analyzed (J, K). A large proportion of the caspase-3+ cells were also BrdU+ (black line). Each of the graphs is a two-point average plot of one individual for each survival time.
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axis, we analyzed consecutive transverse sections along a 1cm segment of the regenerating spinal cord at post-injection intervals ranging between 2 h and 200 days (n = 22 fish). The analyzed region always included the complete regenerate, plus an intact portion of the spinal cord and the interface between the two. In all analyzed fish from 2 h up to 100 days post-injection survival (n = 18), a peak in the number of BrdU+ cells and another peak in the number of active caspase-3+ cells could be observed along the longitudinal axis (Figs. 4A–I). At 150 and 200 days post-injection, the peak of apoptotic cells disappeared, whereas the peak of BrdU+ cells persisted (Figs. 4J, K). Although the absolute magnitude of these peaks varied between time points and individuals, a similar pattern was evident in all cases. At short post-injection survival times (up to 15 days post-amputation/5 days post-BrdU injection), the high density of BrdU+ cells started at, or immediately next to, the lesion site, extending up to approximately 1000 μm rostrally. A peak of apoptotic cells, usually much smaller in magnitude, followed at approximately 500–1500 μm rostral to the lesion site. Numerous apoptotic cells were double-labeled. Interestingly, at later time points, the cell proliferation peak could be observed increasingly farther from the caudal tip of the regenerating spinal cord, up to 4000–6000 μm at 100– 200 days post-injection (Figs. 4I–K). Since the spinal cord continues to grow caudally, the fact that the BrdU peak remained relatively stationary indicated that few of the dividing cells labeled at the time of the BrdU injection migrated caudally over long distances. The apoptosis peak was always seen rostral to the proliferation peak, which was very close to the initial injury site. Fig. 5 schematically shows the regrowth of the spinal cord and the distribution of caspase-3+, BrdU+, and double-labeled cells relative to the site of the lesion.
2.4. cells
Apoptosis after differentiation of newly generated
To examine the relationship between apoptotic cells and differentiated cells among the newly generated ones, we combined immunolabeling against active caspase-3 and BrdU with labeling against the early neuronal marker Hu (Marusich and Weston, 1992) or the astroglial marker GFAP (Eng et al., 2000). Examples of such labeling are shown in Figs. 6 and 7.
2.4.1.
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Fig. 5 – Tail amputation paradigm and spinal cord regeneration after the lesion. (A) A. leptorhynchus with the location of the spinal cord (dotted line). The site where the tail was amputated is indicated by the dashed line. Ten days after the amputation, a single dose of BrdU was administered through intraperitoneal injection, and the fish were killed at various post-BrdU-administration survival times. (B) Schematic representation of the regenerating spinal cord. BrdU application labels numerous proliferating cells in the vicinity of the lesion at all post-amputation survival times (red dots). The majority of BrdU+ cells remain localized at the site of origin, but a few migrate caudally into newly generated tissue. Apoptotic caspase-3+ cells (green dots) are located mostly at the site of the injury, where they persist at levels above-baseline up to at least 110 days after the injury. At the lesion site, most of the caspase-3+ cells are also BrdU+ (black dots), indicating that most of the cells that undergo apoptosis at this location are newly generated ones. Baseline levels of caspase-3+ and BrdU+ cells can be observed in the intact portion of the spinal cord (shown to the right of the transection site). The dashed line indicates the site of transection. Bars = 1 cm in A, 1 mm in B.
Active-caspase-3/BrdU/Hu triple labeling
Immunolabeling confirmed the presence of large numbers of newly differentiated BrdU+/Hu+ neurons in the regenerating A. leptorhynchus spinal cord. Some of these new neuronal cells underwent apoptosis, as indicated by triple caspase-3+/ BrdU+/Hu+ immunostaining (Fig. 6). To quantitatively analyze the occurrence of apoptosis among the cell types, we determined the total number of caspase-3+ cells in spinal cord sections from fish treated as described above, while checking each such cell for double- or triple-labeling against BrdU, Hu, or both (Fig. 8A). This analysis indicated that neurons died at relatively constant rates (0.5 ± SEM: 0.04 cells per section) up to 100 days post-injection, comprising approximately 38% (±SEM: 3%) of apoptotic cells. At 150 days, this value decreased to 0.09 cells
per section (25% of apoptotic cells), while at 200 days only 0.006 cells per section (2% of dying cells) expressed neuronal markers. Triple-labeled caspase-3+/BrdU+/Hu+ cells were observed, to various degrees, at all survival times except at 200 days (Fig. 8A). While at 2 h post-injection only 0.14 cells per section were triple-labeled (8% of the total number of apoptotic cells), this value reached 0.3 cells per section (16% of apoptotic cells) at 1–3 days, and remained relatively constant at 0.3 ±SEM: 0.03 cells per section (25% ±SEM: 2% of apoptotic cells) between 5 and 100 days post-injection. At 150 days post-injection triple-labeled cells amounted to only 0.04 cells per section (10% of apoptotic cells), while at 200 days post-injection no triple-labeled cells could be observed. It
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Fig. 6 – Apoptotic cell death of newly differentiated neurons in the regenerated spinal cord 40 days post-amputation and 30 days post-BrdU administration. (A–D) Triple immunolabeling against active caspase-3, BrdU, and the neuronal marker Hu C/D revealed that some of the newly generated neurons (BrdU+/Hu+; short arrows) are also caspase-3+ (filled arrowheads). Such triple-labeled cells were often found at peripheral locations within spinal cord sections. Apoptotic neurons that were not BrdU+ were also observed (open arrowheads). Red and green lines in D indicate the level of the z-stack cross-section shown on the sides of the panel. The section was taken from the regenerated part of the spinal cord. The central canal is indicated by c. Bar = 100 μm.
should be noted that, due to the administration of a single dose of BrdU after amputation, it is evident that the absolute numbers of new neuronal cells and, consequently, of apoptotic new neurons, are underestimated in the results shown here.
2.4.2.
Active-caspase-3/BrdU/GFAP triple labeling
Similarly to neurons, a certain portion of the newly generated BrdU+/GFAP+ astrocytes died via apoptosis in the regenerating spinal cord (Fig. 7). When analyzed quantitatively, caspase-3+/ GFAP+ astroglia levels remained relatively constant at 0.3 ± SEM: 0.05 cells per section, (24% ±SEM: 1% of apoptotic cells) at any of the analyzed time points, indicating the existence of a definite baseline level of astrocytic apoptosis in the regenerating spinal cord (Fig. 8B). Triple-labeled caspase-3+/BrdU+/GFAP+ cells were observed in very low numbers as early as 2 h after a BrdU injection, when they reached only 0.03 cells per section (1% of all apoptotic cells). The numbers of triple-labeled cells
increased to 0.18 ± SEM: 0.015 cells per section (14% ±SEM: 1.5% of apoptotic cells) between 1 and 20 days post-injection, and then decreased slightly to approximately 0.12 ± SEM: 0.04 cells per section (9% ± SEM: 2.6% of apoptotic cells) between 30 and 100 days. At 150 days, only 0.01 cells per section were triple-labeled (3% of the apoptotic cells), while at 200 days no such cells could be found anymore, further confirming a return to baseline levels in terms of differentiation and integration of new cells (Fig. 8B).
3.
Discussion
3.1. Apoptosis: regulatory function during spinal cord regeneration? Previous work has demonstrated the enormous potential of A. leptorhynchus as a model of successful regeneration of the adult CNS after injury (for reviews, see Zupanc 2008a,b, 2009).
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Fig. 7 – Triple immunolabeling against caspase-3, BrdU, and the astroglial marker GFAP 40 days post-amputation and 30 days post-BrdU administration. (A–D) New astrocytes (BrdU+/GFAP+) were observed throughout spinal cord sections (arrows). Some of these new cells were also caspase-3+ (filled arrowheads). Apoptotic astrocytes that were not BrdU+ were also observed (A, C, D, open arrowheads). Red and green lines in D indicate the level of the z-stack cross-section shown on the sides of the panel. The section was taken from the regenerated part of the spinal cord. The central canal is indicated by c. Bar = 100 μm.
This ability has been documented in both the brain and the spinal cord. For example, a stab wound lesion in the cerebellum of this species has been shown to heal within a few weeks (Zupanc et al., 1998; Zupanc and Ott, 1999). Similarly, within 5 weeks after amputation of the caudal portion of the spinal cord the fish show complete structural and functional regeneration (Sîrbulescu et al., 2009). This high regenerative potential is made possible by massive cell proliferation induced by the injury, and by successful differentiation and integration of the new cells into the existing neural network (Zupanc and Ott, 1999; Sîrbulescu et al., 2009). A second factor that may facilitate wound healing is apoptosis. This type of cell death is likely to minimize tissue inflammation during elimination of damaged cells at the lesion site (Zupanc et al., 1998; Sîrbulescu et al., 2009). Furthermore, the results of the present study are also consistent with the notion that apoptotic cell death might be involved in the development of new tissue. A functional role of apoptosis during the formation of new tissue is supported by observations in a variety of organisms,
ranging from invertebrates to mammals. For example, apoptosis has been shown to act as a mechanism for regulating neuronal connectivity (for reviews, see Chan et al., 2002; Buss et al., 2006). Moreover, in adult organisms in which cell proliferation persists either throughout the CNS or in restricted areas, apoptosis appears to ensure that supernumerary cells are eliminated (Buss et al., 2006). After amputation of the caudal spinal cord of A. leptorhynchus, complete re-growth of nervous tissue occurs, as it does during normal development. It is, therefore, likely that apoptosis after the initial cell elimination phase also plays a role in the structuring of new tissue and in ensuring proper integration of the new cells. In support of this notion, we have found in the present study that up to 110 days post-amputation approximately 38% of the observed apoptotic cells were differentiated neurons, while approximately 24% were GFAP+ astrocytes. These results indicate that a large portion of cells die during later stages of development, presumably at their target areas. This is in agreement with earlier findings in the cerebellum of A. leptorhynchus, where approximately 50% of the newly
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Fig. 8 – Quantitative analysis of caspase-3+ cells in the caudalmost 1 cm of spinal cord after triple-immunolabeling against caspase-3, BrdU, and Hu C/D (A) or GFAP (B). BrdU was administered intraperitoneally 10 days after tail amputation. (A) Analysis indicated that up to 100 days post-BrdU administration survival, neurons died at a relatively constant rate, comprising on average 38% (±SEM: 3%) of apoptotic cells. Newly differentiated neurons (caspase-3+/BrdU+/Hu+) constituted 8% of apoptotic cells at 2 h post-BrdU administration, increased to 16% at 1–3 days, and then maintained a relatively constant level of approximately 25% (±SEM: 2%) of apoptotic cells between 5 and 100 days post-BrdU administration. After 100 days, these percentages decreased, with neurons representing only 2% of apoptotic cells. No apoptotic new neurons were observed at 200 days. (B) Astrocytes constituted on average 24% (±SEM: 1%) of apoptotic cells at all time points analyzed. Apoptotic newly differentiated astroglia (caspase-3+/BrdU+/GFAP+) were present at very low levels (1%) at 2 h post-injection. Their numbers increased to 14% (±SEM: 1.5%) between 1 and 20 days, followed by a decline to approximately 9% (±SEM: 2.6%) between 30 and 100 days, and to 3% by 150 days post-injection. At 200 days post-injection no triple-labeled cells could be observed.
generated cells die upon arrival at the target sites, presumably by undergoing apoptosis (Soutschek and Zupanc, 1996; Ott et al., 1997). Is there a selection process that determines which cells undergo apoptosis? A previous study in the intact brain of A.
leptorhynchus has shown that apoptosis affects preferentially those adult-born cells that exhibit numerical chromosome aberrations due to mitotic segregation defects (Rajendran et al., 2008). It is likely that aneuploidy also occurs among the new cells generated in response to injury, because they derive from the same pool of stem cells as the new cells in the intact brain (Zupanc and Ott, 1999). It would, therefore, be interesting to examine whether part of the new cells generated after spinal cord injury are also aneuploid, and if so, whether these cells are eliminated with higher frequency through apoptosis than euploid cells. Interestingly, we observed an uneven radial distribution of newly generated cells undergoing apoptosis within the regenerating spinal cord. Such cells were present in relatively low numbers in the proximity of the central canal and the pial surface (Fig. 2B), suggesting a protective effect in these areas. In other regeneration-competent organisms, such as urodele amphibians, ependymal cells have been shown to upregulate trophic factors, such as retinoic acid and fibroblast growth factor, after spinal cord lesion (Chernoff et al., 2003; Ferretti et al., 2003). Furthermore, fibroblast growth factor has been demonstrated to effectively inhibit caspase-3 activation in vitro (Miho et al., 1999). Since an ependymal response to injury also occurs in A. leptorhynchus, it is possible that local diffusion of growth factors might inhibit apoptosis in the proximity of the central canal. Why apoptosis levels are relatively low close to the pial surface remains unclear, but perhaps the glia limitans exerts a similar trophic function as do the ependymal cells at the central canal. Another line of evidence in favor of the restructuring function of apoptosis during the cell replacement phase of spinal cord regeneration is the observed caudo-rostral gradient of caspase-3+ cells. While very few caspase-3+ or doublelabeled caspase-3+/BrdU+ cells were found in the caudalmost (young) regions of the regenerated spinal cord, the number of such cells gradually increased, relative to baseline levels, in more rostral (older) parts (Fig. 4). Corroborated by the fact that spinal cord differentiation after tail amputation takes place in a rostro-caudal direction (Waxman and Anderson, 1986; Chernoff et al., 2003; Ferretti et al., 2003), this finding indicates that most apoptotic events do not occur at the site of cell proliferation, but instead in regions of cell maturation. In addition, while apoptotic cells constituted, on average, 15% of the BrdU+ cells in the period of 1–100 days after BrdU administration, between 60 and 90% of apoptotic cells were also BrdU+ during the same interval. This result demonstrates that the majority of apoptotic cells found throughout the analyzed portion of the spinal cord were, indeed, young cells generated after tail amputation. Taken together, these results support the hypothesis that apoptosis plays a regulatory role during nervous tissue regeneration in the spinal cord of A. leptorhynchus, presumably by eliminating superfluous or poorly integrated cells.
3.2. Elevated levels of apoptosis at the transition zone between intact and regenerated tissue After the initial phase of cell elimination by apoptosis, the number of apoptotic cells remained low throughout the intact, as well as the regenerated parts of the spinal cord, with one
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exception. At the site where the spinal cord had been cut, a pronounced peak of apoptotic cells, sometimes three times higher than baseline levels, could be observed up to 110 days after amputation (Fig. 4). Such long-term persistence of a relatively high frequency of occurrence of apoptotic cell death at this specific location may reflect a defective, or incomplete, organization of the transition zone between the intact tissue and the newly generated one. This finding is especially remarkable since the regenerated part of the spinal cord does receive innervation from intact regions (Sîrbulescu et al., 2009). It is possible that the damage inflicted by the lesion induces a long-lasting alteration of the local environment at the interface between the intact part and the regenerate, which significantly delays successful integration, but not differentiation, of new cells. The distribution of triple-labeled caspase-3+/BrdU+/Hu+ and caspase-3+/BrdU+/ GFAP+ cells at this location mirrored the distribution of caspase-3+ cells, indicating that the observed apoptotic peak included differentiated neurons and glia, and could be located at an integration interface within the regenerating spinal cord. Interestingly, a previous study in a related teleost species, which is capable of regenerating its tail after amputation similarly to A. leptorhynchus, has also revealed structural modifications at the transition zone between regenerated and unregenerated spinal cord (Anderson et al., 1984). In the latter study, longitudinal sections through spinal cord have shown a clear constriction of the tissue around the transition area, 4 months after amputation. Albeit not of the same magnitude, we have also observed some decrease in tissue density in transverse spinal cord sections at this level in A. leptorhynchus, suggesting that ongoing apoptosis exceeds cell proliferation in this area, and thus affects the structure of the tissue. In the present study, we monitored apoptotic cell death starting 10 days post-lesion. At this time point, the number of apoptotic cells at the site of injury was only one tenth of the levels found immediately after the amputation when massive elimination of cells through apoptosis occurs (Sîrbulescu et al., 2009). Furthermore, the peak of apoptosis regressed after 110 days post-amputation and was not visible anymore at 160 or 210 days after the lesion (Fig. 4). These results indicate that the long-lasting disruption of tissue at the original site of the cut is eventually resolved, and successful integration is finally achieved. At the same time, 150–200 days post-BrdU administration, the proportion of newly differentiated neurons and astroglia that undergo apoptosis becomes markedly lower (Fig. 8). This might represent the end of the integration process of the new cells, and the return of the tissue to a cytoarchitecture reminiscent of the pre-injury structure.
3.3.
New roles of apoptosis: a comparative view
In mammals, apoptotic cell death takes place not only during the acute phase associated with the primary injury, but also during secondary injury which extends for long periods of time after the initial insult (for reviews, see Beattie et al., 2002; Kwon et al., 2004; Profyris et al., 2004; Fitch and Silver, 2008; Johansson, 2007; Ling and Liu, 2007). Elevated levels of apoptosis during the secondary injury can last for many
23
weeks after the lesion (Beattie et al., 2002; Kwon et al., 2004; Keane et al., 2006; Bramlett and Dietrich, 2007). Such high levels of cell death are considered one of the main reasons for the significant loss of neural tissue after spinal cord injury (Kim et al., 2003; Martiñón and Ibarra, 2008), thus seemingly supporting the notion of a detrimental role of apoptosis after injury in non-regenerating systems. By contrast, in regeneration-competent systems there is evidence of a rather supportive function of apoptosis after injury. During the first few days after experimentally applied lesions, apoptosis appears to mediate the “clean” elimination of damaged cells in both the brain (Zupanc et al., 1998) and the spinal cord of A. leptorhynchus (Sîrbulescu et al., 2009). As suggested by the present study, apoptosis might also be involved in tissue repair during the later stages of spinal cord regeneration by regulating the survival of newly formed cells. This is consistent with the observation in larval lampreys where apoptotic cell death persists for at least 12–16 weeks after spinal cord transection (Shifman et al., 2008). In Xenopus laevis tadpoles, inhibition of apoptosis has been shown to effectively prevent regeneration after tail transection (Tseng et al., 2007). Interestingly, in rats recent evidence indicates that preventing apoptosis for an extended duration after spinal cord injury may impair recovery and lead to an increase in necrosis and inflammation (Cittelly et al., 2008). These studies suggest that apoptosis does not only exert a negative function after a lesion by eliminating cells, as traditionally assumed. Instead, it may also play an important role for the development of new cells that replace tissue lost to injury. The spinal cord of A. leptorhynchus will provide an excellent regeneration-competent model system to test this hypothesis.
4.
Experimental procedures
4.1.
Animals
Fish were purchased from a tropical fish importer and kept in individual aquaria under defined conditions (Zupanc et al., 2006). A total of 25 adult fish with a total body length between 102 mm and 154 mm (mean: 126 mm; median: 126 mm) and body mass between 1.9 g and 8.2 g (mean: 4.1 g; median: 4.5 g) were used in this study. Sixteen fish were males and nine females, as revealed by post-mortem gonadal inspection. The gonadosomatic index, determined as fresh weight of gonads divided by body mass, ranged between 0.0013 and 0.005 (mean: 0.0032; median: 0.0030) in males, and between 0.0068 and 0.1841 (mean: 0.0904; median: 0.0940) in females. All experiments were carried out in accordance with the relevant German law, the Deutsches Tierschutzgesetz of 1998.
4.2.
Immunohistochemistry
Under general anesthesia with urethane (Sigma, Taufkirchen, Germany) and after application of 2% lidocaine (Euro OTC Pharma, Hannover, Germany) as a local anesthetic, 1 cm of the tail, measured rostrally from the white caudal ring, was amputated (n = 25 fish). After a 10-day recovery period, the
24
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fish received a single intraperitoneal injection of approximately 50 μl of a 3 mg/ml BrdU solution (Amersham/GE Healthcare, Freiburg, Germany). Following various post-amputation survival times (2 h, 1 day, 3 days, 5 days, 10 days, 20 days, 30 days, 40 days, 50 days, 100 days, 150 days, 200 days; n = 2 fish per survival time), animals were deeply anesthetized using ethyl 3-aminobenzoate methanesulfonate (MS-222; Sigma) and intracardially perfused. The 1-cm caudalmost segment was removed from the tail, post-fixed, and cryoprotected as described (Zupanc et al., 2005). Transverse or sagittal sections were cut at a thickness of 16 μm and mounted serially onto SuperFrost Plus Gold slides (Menzel-Gläser, Braunschweig, Germany).
4.2.1.
Active caspase-3/BrdU double immunolabeling
After the sections had been dried in a desiccator for 90 min at room temperature (RT), they were rinsed three times for 10 min each in 0.1 M tris-buffered saline (TBS) and blocked for 60 min with 3% normal sheep serum, 1% teleostean gelatin, 1% bovine serum albumin (BSA), and 0.3% Triton X-100 in TBS (all from Sigma). Sections were incubated with a purified monoclonal rabbit anti-active caspase-3 antibody (clone C92-605; Cat. No. 559565; BD Pharmigen, Heidelberg, Germany), diluted 1:100 in blocking solution, at 4 °C overnight. After three washes in TBS for 5 min each, the sections were blocked for 30 min at RT with 3% goat serum (Sigma), 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100, followed by incubation for 90 min at RT with the secondary antibody Alexa Fluor 488conjugated goat anti-rabbit IgG (Molecular Probes/Invitrogen, Karlsruhe, Germany), diluted 1:200 in blocking solution. After three washes in TBS for 10 min each, the sections were first incubated with a solution of 20% acetic acid and 70% ethanol in ddH2O for 5 min at RT, and then in 70% ethanol for 25 min at −20 °C. After three washes in TBS for 5 min each, the sections were incubated in 2 M HCl at 37 °C for 30 min, followed first by two rinses in 0.1 M borate buffer (pH 8.5) at RT for 20 min each, and then by six rinses in TBS for 5 min each. After blocking for 30 min with 3% normal sheep serum, 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100 in TBS, the sections were incubated with rat anti-BrdU antibody (Cat. No. OBT0030; Oxford Biotechnology through Biozol, Eching, Germany), diluted 1:200 in blocking solution, at 4 °C overnight. Following three rinses in TBS for 5 min each, the sections were blocked for 30 min at RT with 3% donkey serum (Sigma), 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100, and incubated for 90 min at RT with the secondary antibody, donkey anti-rat IgG conjugated to Cy3 (Jackson Immunoresearch through Dianova, Hamburg, Germany), at a dilution of 1:1500 in blocking solution. After three washes in TBS for 10 min each, the sections were embedded in polyvinyl alcohol containing n-propyl gallate.
4.2.2.
Active caspase-3/BrdU/Hu C/D triple immunolabeling
After drying in a desiccator for 90 min at RT, the sections were rinsed three times for 10 min each in 0.1 M phosphatebuffered saline (PBS), incubated for 30 min in 50 mM Tris, pH 8.0, and finally rinsed three times in PBS for 5 min each. Blocking was performed for 60 min with 3% normal sheep serum, 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100 in PBS, after which sections were incubated with a monoclonal
mouse anti-Hu C/D antibody (clone 16A11; Cat. No. A-21271; Molecular Probes), diluted 1:20 in blocking solution, at 4 °C overnight. After three washes in PBS for 5 min each, the sections were blocked for 30 min at RT with 3% goat serum, 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100 in PBS, followed by incubation for 90 min at RT with the secondary antibodies Alexa Fluor 635-conjugated goat anti-mouse IgG (Molecular Probes), diluted 1:200, and Coumarin AMCAconjugated goat anti-mouse IgG (Jackson Immunoresearch), diluted 1:100, in blocking solution. Further labeling for active caspase-3 and BrdU was performed as described above.
4.2.3.
Active caspase-3/BrdU/GFAP triple immunolabeling
After drying in a desiccator for 90 min at RT, the sections were rinsed three times for 10 min each in 0.1 M TBS, and blocked for 60 min with 3% normal sheep serum, 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100 in TBS. Sections were incubated with a monoclonal mouse anti-GFAP antibody (clone G-5-A; Cat. No. G3893; Sigma), diluted 1:50 in blocking solution, at 4 °C overnight. After three washes in TBS for 5 min each, the sections were blocked for 30 min at RT with 3% goat serum, 1% teleostean gelatin, 1% BSA, and 0.3% Triton X-100 in TBS, followed by incubation for 90 min at RT with the secondary antibodies Alexa Fluor 635conjugated goat anti-mouse IgG (Molecular Probes), diluted 1:200, and Coumarin AMCA-conjugated goat anti-mouse IgG (Jackson Immunoresearch), diluted 1:100, in blocking solution. Further labeling for active caspase-3 and BrdU was performed as described above.
4.3.
Antibody controls
Specificity of the rabbit anti-active caspase-3, rat-anti BrdU and mouse anti-GFAP primary antibodies in A. leptorhynchus was demonstrated previously both by pre-adsorption of the respective primary antibody with the specific antigen and by omission of the primary antibody during labeling (Clint and Zupanc, 2001; Sîrbulescu et al., 2009). The mouse anti-Hu C/D antibody used in the present study has previously been shown to recognize a single protein band at the expected molecular weight of 42 kDa in Western blots of tissue from teleostean CNS (Takeda et al., 2008).
4.4.
Microscopy and analysis
Sections were examined under a Zeiss Axioskop epifluorescence microscope (Carl Zeiss, Göttingen, Germany). Every complete section was analyzed, and all caspase-3-positive cells were counted. In double-labeling experiments, also all BrdU-positive cells were counted. In any experiment, a minimum of 64 sections were analyzed per fish. The location of labeled cells was entered into camera lucida drawings of spinal cord sections. Every caspase-3-positive cell was checked for double- or triple-labeling against other antigens. For confocal microscopy, specimens were examined on a Zeiss LSM 510 META laser-scanning microscope. Optical sections were taken with a pinhole opening of 1 Airy and at a resolution of 1024 × 1024 pixels, using LSM 5 (version 3.2; Carl Zeiss) software. For longitudinal sections, multiple consecutive images taken along the available portion of the spinal cord
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were digitally stitched using CorelDraw 11.0 (Corel Corporation, Ottawa, Canada) to give an overview of the reconstructed tissue section.
Acknowledgments This study received financial support from the Ernst A.-C. Lange-Stiftung and from Jacobs University Bremen.
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