Dynamics of degeneration and regeneration in developing zebrafish peripheral axons reveals a requirement for extrinsic cell types
© Villegas et al.; licensee BioMed Central Ltd. 2012
Received: 28 November 2011
Accepted: 1 May 2012
Published: 8 June 2012
Understanding the cellular mechanisms regulating axon degeneration and regeneration is crucial for developing treatments for nerve injury and neurodegenerative disease. In neurons, axon degeneration is distinct from cell body death and often precedes or is associated with the onset of disease symptoms. In the peripheral nervous system of both vertebrates and invertebrates, after degeneration of detached fragments, axons can often regenerate to restore function. Many studies of axonal degeneration and regeneration have used in vitro approaches, but the influence of extrinsic cell types on these processes can only be fully addressed in live animals. Because of its simplicity and superficial location, the larval zebrafish posterior lateral line (pLL) nerve is an ideal model system for live studies of axon degeneration and regeneration.
We used laser axotomy and time-lapse imaging of pLL axons to characterize the roles of leukocytes, Schwann cells and target sensory hair cells in axon degeneration and regeneration in vivo. Immune cells were essential for efficient removal of axonal debris after axotomy. Schwann cells were required for proper fasciculation and pathfinding of regenerating axons to their target cells. Intact target hair cells were not themselves required for regeneration, but chemical ablation of neuromasts caused axons to transiently deviate from their normal paths.
Macrophages, Schwann cells, and target sensory organs are required for distinct aspects of pLL axon degeneration or regeneration in the zebrafish larva. Our work introduces a powerful vertebrate model for analyzing axonal degeneration and regeneration in the living animal and elucidating the role of extrinsic cell types in these processes.
KeywordsIn vivo axotomy Wallerian degeneration Schwann cells Leukocytes Hair cells Neurons Lateral line
Axonal degeneration occurs during normal development of the nervous system and is central to the pathology of neurodegenerative diseases, nerve damage caused by metabolic diseases, and mechanical nerve injuries[1–3]. While there are different types of axonal degeneration, similar mechanisms regulate both developmental pruning of excessive axonal branches and the selective removal of damaged axons[1, 4, 5]. Wallerian degeneration (WD) occurs in axon fragments that are separated from their cell body. WD occurs in a sterotyped and orderly fashion, implying that it is under genetic control, and has been described in the central and peripheral nervous system after trauma, stroke or infection[1, 3]. Immediately after an axon is severed, acute axonal degeneration (AAD) can occur at both ends adjacent to the cut. Following AAD, the detached axon fragment remains intact during a characteristic “lag phase”. Following the lag phase, axons quickly degenerate: the endoplasmic reticulum breaks down, neurofilaments degrade, mitochondria swell and the axon fragments. Finally, in the last phase of WD, axon fragments are removed by phagocytic cells.
In the peripheral nervous system (PNS), Schwann cells and macrophages play important roles throughout the process of WD. Schwann cells decrease the synthesis of myelin lipids during the first 12 hours after axotomy and stop producing myelin proteins within 48 hours. In the absence of macrophages, the process is even more rapid[3, 10], with glial cells removing myelin during the earliest stages of PNS axon degeneration[11–14]. Glial cells also release chemokines and cytokines, some of which are responsible for recruiting macrophages to the site of nerve degeneration in the final phase of myelin removal[15, 16]. After injury, these cells adhere to the basal lamina, enter the nerve, and phagocytose opsonized debris[17–20].
The limited regenerative capacity of neurons in the adult central nervous system (CNS) of mammals has been a subject of intense study. Several studies in vertebrate models have established that, in addition to intrinsic growth programs, extrinsic factors regulate axonal regeneration[21, 22]. For example, inhibitory molecules associated with myelin and glial scars are induced by axotomy and create obstacles to axonal regeneration in the CNS. Astrocytes, which form glial scars in the CNS after injury, are absent in the peripheral nervous system (PNS), though inhibitory molecules associated with myelin are expressed in the PNS. However, in the PNS, Schwann cells undergo dedifferentiation after injury, diminishing the effect of inhibitory proteins. Schwann cells and macrophages in the PNS can have positive effects on regeneration by rapidly removing myelin debris and expressing a wide range of neurotrophic factors that create a favorable environment for axonal growth. Glia can also serve as guides by providing structural substrates to facilitate the growth of axons along their original paths.
The zebrafish lateral line (LL) has been a useful model for understanding interactions between axons and extrinsic cell types, including glial cells, during development, but it has yet to be exploited for studies of axon degeneration and regeneration. The LL is a mechanosensory system that responds to mechanical stimuli produced by water movement. It is composed of groups of individual sensory organs called neuromasts, which are distributed on the body surface in species-specific patterns. Each neuromast contains 15 to 20 hair cells at its core, surrounded by two types of accessory cells: supporting cells and mantle cells. Hair cells are innervated by afferent sensory neurons whose cell bodies are located in either the anterior or posterior LL ganglia. LL neurons extend their central axons to the hindbrain[29–31] in a somatotopic fashion[32, 33]. Approximately 20 bipolar afferent neurons receive synaptic input from hair cells of each major branch of the lateral line. Most studies to date have focused on the posterior LL (pLL), which extends along the trunk and tail.
To determine whether extrinsic cell types can influence axon degeneration or regeneration in the PNS, we have characterized these processes after pLL nerve axotomy. This approach allowed us to quantitatively describe the onset and progression of WD in axotomized pLL neurons and to follow their regeneration. We found that removal of glia, leukocytes, and target cells each had a distinct effect on different aspects of axon degeneration or regeneration. Together, these studies establish the zebrafish pLL nerve as a powerful model for live studies of axon degeneration and regeneration and uncover a rich variety of cell-cell interactions that regulate these processes.
Lateral line axons undergo Wallerian degeneration after axotomy
The zebrafish posterior lateral line (pLL) is an excellent model for studying peripheral axonal structure and function in vivo. The pLL nerve is long and superficially located, target cells in neuromasts are located along the body surface in stereotyped positions, and all cell types in the system can be genetically, physically or chemically ablated. These properties made it possible for us to use laser axotomy and time-lapse imaging to monitor axon degeneration and regeneration after injury to lateral line axons in live zebrafish larvae. To study the behavior of the entire nerve we used the neuroD::EGFP stable transgenic line, and to analyze the behavior of single neurons we injected the HuC::GFP transgene at the single cell stage and screened for transient transgenic embryos expressing GFP in single lateral line neurons at three days postfertilization (dpf). Thus, we were able to transect all axons in the nerve using the NeuroD::EGFP transgenic fish line, and sever single axons with HuC::GFP transient transgenics, presumably leaving the rest of the pLL nerve intact. Neurons were axotomized at 78 hours postfertilization (hpf) using a two-photon microscope and imaged at one- two- or twenty-minute intervals for up to twelve hours with confocal microscopy. We chose three dpf fish because at this stage the pLL system and innate immune leukocytes are functional. At three dpf Schwann cells have differentiated, overlie the pLL nerve, and express myelin, though the myelin sheath only forms later, between four and seven dpf[35, 36].
To determine whether fragmentation occurs progressively as a wave from the lesion site to the axon terminal (proximal to distal), as has been described in other systems[7, 38–40], synchronously along the entire axon, or distal to proximal, we imaged single severed axons every two minutes after axotomy at two different positions along the axon fragment: adjacent to the injury site and approximately 800 to 1000 μm further down the axon (Additional Files3,4 and5). Imaging fragmentation at these two regions revealed that the most distal region fragmented before the more proximal region (n = 6; Additional Files3 and4). Imaging shorter (500 μm) fragments indicated that the timing and progression of fragmentation varied widely from axon to axon (Additional File5). In some cases, fragmentation occurred synchronously along the fragment (Additional File5A) and, in others, advanced in a ‘saltatory’ fashion (Additional File5B).
Schwann cells regulate the number of axon fragments produced during Wallerian degeneration of the posterior lateral line nerve
Leukocytes regulate the onset of axon fragment clearance during Wallerian degeneration of the posterior lateral line nerve
To test the hypothesis that leukocytes mediate phagocytosis, we used a morpholino targeting the spi1 gene, which specifically ablates the myeloid lineage (see Materials and Methods). Comparing WD kinetics in these fish and controls indicated that the timing of axon fragmentation was normal but that the clearance phase was significantly longer in immune cell-depleted fish (Figure2), as axon fragments persisted for over six hours under these conditions (Figure3). When we combined immune cell and Schwann cell depletion, the speed of clearance was restored to near control levels (Figure2), indicating that cell types other than innate immune leukocytes can contribute to debris clearance, but only when Schwann cells are absent. Epidermal skin cells are strong candidates, since lateral line axons interact closely with skin cells in the absence of glia and have been suggested as potential phagocytes of degenerating trigeminal axon fragments in zebrafish. Double mutant fish lacking both leukocytes and Schwann cells did not survive long enough after axotomy for regeneration to be examined (data not shown).
Posterior lateral line innervation does not influence survival, differentiation, or regeneration of hair cells
Lateral line axons regenerate along their original trajectories
Schwann cells are required for proper regeneration
Axonal regeneration does not depend on intact target hair cells but is negatively affected by copper-induced ablation of neuromasts
To characterize the influence of nonautonomous tissues on axon degeneration and regeneration, we developed a reproducible method for severing the zebrafish pLL nerve in vivo and monitoring its behavior with time-lapse imaging. Imaging allowed us to visualize disconnected pLL axon fragments undergoing Wallerian degeneration and to monitor the entire process of axon regeneration, from injury to the reestablishment of pLL functionality. This system thus offers an unprecedented opportunity for analyzing the regulation of axon degeneration and regeneration in vivo and dissecting the roles of cellular and molecular regulators of these processes.
WD in the zebrafish pLL was extremely fast compared to axon degeneration in many previously characterized models. For example, the lag phase in the dorsal root ganglion nerve of the mouse lasts one to one and a half days after axotomy, a phenomenon that in the pLL of zebrafish larvae takes approximately three hours, similar to what was recently described for spinal motor neurons. The elimination of axonal fragments took an average of around five hours in the pLL and twenty-four hours in spinal motor neurons. Both of these phases are slightly but reproducibly faster in zebrafish trigeminal axons. This observation indicates that WD in peripheral axons differ even in the same animal, suggesting either intrinsic or extrinsic differences between them. The fact that we were able to assess the success of WD just five to six hours after axotomy raises the possibility that medium-to-high throughput strategies for drug assays or genetic screens may be a feasible method for dissecting WD in this system. Imaging fragmentation along pLL axons (which can be longer than 1000 μm) demonstrated that WD proceeds from the distal end of an axon fragment to the proximal end, even though in a given section of a detached axon, fragmentation can appear to be synchronous. Surprisingly, this retrograde progression more closely resembles the outcome of a crush injury to the mouse sciatic nerve than the anterograde progression seen when the sciatic nerve is severed. The signals mediating propagation along a severed axon fragment in one direction or the other are still unknown, but our results provide an additional model for investigating retrograde progression.
Zebrafish larvae can regenerate many tissues and structures, including the pLL nerve. By performing time-lapse imaging of regenerating axons, we observed new growth cones arising from the proximal axon stump immediately after degeneration of the distal fragment. In both pLL and trigeminal axons, regeneration starts only after debris from the original axon is cleared. It is still not clear whether regeneration is dependent on fragment removal per se since, in trigeminal neurons, regeneration occurs on schedule when axon degeneration is delayed by WldS expression. Nonetheless, these neurons seem to have evolved so that degeneration and regeneration occur in orderly succession.
Regeneration of the pLL nerve followed its original pathway, similar to what has been described for regeneration of zebrafish spinal motor neurons. Analyzing regeneration of single axons labeled in transient transgenic larvae, we verified that reconstitution of a functional nerve corresponded to bona fide axon regeneration, rather than axon growth arising from new neurons. Regeneration of pLL axons qualitatively resembled axon regeneration described in invertebrates and in some other vertebrate axons[60, 61]. Regenerating pLL axons in wild-type animals grew at a speed of approximately 0.5 μm/minute, reaching all of the pLL neuromasts by 24 hours postaxotomy. These axons thus regenerated much faster than regenerating axons in the mouse central and peripheral nervous systems, where axons grow at 0.2 μm/minute and 0.07 μm/minute, respectively.
Axon fragments are known to be phagocytosed by at least two cell types: Schwann cells and immune cells. Ample evidence indicates that in the mouse PNS macrophages and glial cells jointly regulate the clearance of axons and myelin[13–16]. We found that in the absence of innate immune cells, pLL axon fragments persisted for significantly longer than in controls (Figure2). Elimination of each cell type separately suggests that leukocytes are the main participants in the removal of axonal debris. Our findings are complementary to those of Rosenberg et al., whose work suggested that debris clearance after motor axon transection is mediated by macrophages, and that Schwann cells are dispensable for this process. We also suggest that at least one other cell type must have the capacity to remove axonal debris because in the absence of both leukocytes and Schwann cells, axon fragments disappeared on schedule. Epidermal skin cells could fulfill this role as they associate closely with the pioneering pLL axons during their development and, in the absence of Schwann cells, pLL axons remain within the epidermis.
Interestingly, we found that Schwann cells are essential for proper navigation of the regenerating pLL nerve. In the absence of these cells, axons grew erratically, even though they extended at normal speeds. In the mammalian inner ear, several studies indicate that hair cells secrete factors that induce axon growth of acoustic ganglion neurons after injury (reviewed in[22, 26]). Moreover, after Wallerian degeneration, the endoneurium that had surrounded the distal axon remains, leaving an endoneurial tube around which Schwann cells proliferate, forming bands of Büngner[61, 62] that serve as a structural guide by which new axons navigate. In addition to this structural role, after nerve injury Schwann cells express neurotrophic factors, which could contribute to axon guidance. The erratic growth observed in regenerating pLL axons in the absence of Schwann cells, may thus be explained by the absence of a structural guide for regeneration or the lack of appropriate guidance molecules.
A recent report identified a fundamental role for glial cell line-derived growth factor (GDNF) in axonal guidance during development and regeneration of the pLL nerve. In contrast with our results, that study found that Schwann cell absence did not affect regeneration of the pLL nerve, but that regeneration was dependent on GDNF production by interneuromastic cells. However, in that report, only a subset of Schwann cells were ablated (specifically, Schwann cells between neuromasts I and II). It is possible that some Schwann cells escaped ablation (for example, by dedifferentiating and losing GFP transgene expression) or that intact Schwann cells located beyond the second neuromast continued to exert an effect on the regenerating nerve.
Our study also addressed whether neuromast hair cells, the targets of innervation by the pLL nerve, are necessary for proper regeneration or pathfinding of axons in the pLL. Chemical elimination of neuromast cells (using copper toxicity) resulted in erratic growth of regenerating pLL axons, but axons reestablished the correct pathway upon regeneration of the neuromasts. This result suggests that the target cell environment influences the guidance of regenerating axons. However, when hair cells were eliminated genetically or physically with a laser, pLL regeneration trajectory and velocity were both normal, suggesting that hair cells themselves are not required for this process. A possible explanation for these contradictory results is that although copper treatment selectively damages neuromasts, without affecting surrounding glial cells, axons, or other tissues in the pLL, it can cause damage to multiple cell types in the neuromast, such as supporting cells and mantle cells, in addition to hair cells. These other cell types in the neuromast were not affected by laser or genetic ablation. Thus, it is possible that the aberrant behavior of regenerating pLL neurites observed after copper treatment is due to neuromast accessory cell elimination. We cannot, however, exclude the possibility that copper may have an unexpected effect on the regenerating axons themselves or on the interstitial space through which they navigate. It will be important to use alternative methods of ablating neuromasts to further test their role in pLL axon regeneration.
While we did not find that innervation was required for the survival of most hair cells, it is possible that viability of these mechanosensory cells would be impaired after long-term denervation by afferent fibers, as is clearly the case in the mammalian inner ear. We observed occasional hair cell death after axotomy, but most dying cells identified by TUNEL staining were located among accessory cells in denervated neuromasts. While it was known that pLL neuromast hair cells can develop and differentiate in the absence of a pLL nerve[47, 48], this is the first study to examine their viability when denervation occurs postembryonically, after hair cells have developed and integrated normally into the lateral line. Nonetheless, reinnervation occurred within 24 hours after injury, due to the robust regeneration of the proximal axon, indicating that the hair cells are deprived of afferents for only a short period. It would be interesting to permanently block axonal regrowth and assess the survival of hair cells in the denervated state.
The model we have developed exploits the high degree of conservation among the elements of the mechanosensory systems of vertebrates. The mechanisms of nerve degeneration, including WD, can be studied in a straightforward manner in the zebrafish pLL, offering a new tool to identify molecules and genes that regulate axon degeneration and regeneration. In this work, we have provided evidence for the roles of extrinsic cell types in the processes of lateral line axon degeneration and its subsequent regeneration, roles which are distinct and necessary for reestablishing a functional circuitry in this organ.
Zebrafish husbandry and genetic strains
Tübingen and AB wild-type, transgenic and mutant strains of zebrafish (Danio rerio) were maintained at 28.5°C on a 14-hour light/10-hour dark cycle. The transgenic strains used were: neuroD::EGFP kindly provided by Dr. Alex Nechiporuk, brn3c::mGFP from Dr. Herwig Baier, lysC::GFP from Dr. Phil Crosier, foxD3::GFP from Dr. Darren Gilmour and mpeg1::mCherry from Dr. Graham Lieschke. We obtained the leo1 mutant from Dr. Jau-Nian Chen.
All embryos were collected by natural spawning, staged according to Kimmel et al. (1995), and raised at 28°C in E3 medium (5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.3 mM MgSO4, and 0.1% methylene blue) in Petri dishes, unless otherwise noted. We express the larval ages in hours postfertilization (hpf) or days postfertilization (dpf). All experiments were carried out in 72 to 78 hpf larvae since, at this stage, the primary lateral line is completely developed and functional.
Antisense morpholino and DNA injection
List of morpholinos used
Approximately 1 ng of HuC::GFP DNA (construct obtained from Dr. Hernán López-Schier) was injected into one-cell-stage embryos to visualize GFP in single lateral line neurons in mosaic larvae. GFP-expressing embryos were selected and axotomized as described below.
Imaging and axotomy
pLL axons were imaged in stable transgenic neuroD::GFP zebrafish larvae and in transient transgenic HuC::GFP larvae. In most experiments, the entire nerve was severed in stable neuroD::EGFP transgenic larvae. A single labeled axon was cut in the case of transient HuC::GFP transgenic fish (for example, Figure1 and Additional Files1,3,4 and5). Embryos were dechorionated, anaesthetized in 0.01% tricaine, and mounted in a sealed agarose chamber. A Zeiss LSM510 confocal/two-photon microscope (Carl Zeiss AG, Oberkochen, Germany) was used to image and axotomize GFP-labeled lateral line neurons. Axons were visualized using a 25x water objective (NA 0.8), a laser wavelength setting of 910 nm, and 30 mW of power at the sample. To axotomize a specified region of the axon, a single 2D scan with 180 mW at a 70x ScanImage zoom was used. For time-lapse analysis, embryos were imaged at various intervals for one to twelve hours on a confocal microscope with a 20x, 0.5 NA air objective (Zeiss LSM 510). Approximately 15 optical sections were obtained at each time point, spaced 3 μm apart. These were compiled into Z-projections and movies with ImageJ and QuickTime software. Embryos were maintained at 28.5°C with a heated stage throughout imaging. Imaging for longer times or with older stages was not possible since after axotomy and treatments larvae became deformed, necrotic, or died. mpeg1::cherry transgenic larvae (Additional File16) were imaged with an Olympus, MVX10 fluorescence dissecting scope (Olympus, Tokyo, Japan), equipped with a QImaging camera and Micropublisher 3.3, and were analyzed with ImageJ software.
Drugs and copper treatment
Morphological analysis and data quantification
ImageJ software was used to quantify axonal fragments. The confocal images were binarized such that pixel intensity of regions corresponding to axons was converted to black and all other regions were converted to white. Axonal fragments were quantified using the ImageJ Analyze Particles plugin, with the following settings: size pixel ^ 1 = 0-Infinity; circularity = 0–1; show outlines. The times obtained for different phases of WD were determined by blinded visual inspection of stacks and then averaged for each treatment.
Statistics and analysis
We used one-way ANOVA for treatment comparison of parametric data or an equivalent nonparametric method (Kruskal-Wallis). Additionally a two-way ANOVA was used when the parameter depended on two factors (see text for details). The significance level was P <0.05 for all treatments. Values obtained from quantification of fragments were plotted against time. We used the Student’s t test for comparison of spi1 morphants and controls. All data analysis was performed using Prism 5.0 (GraphPad Prism Software, Inc., San Diego, CA, USA).
Rosario Villegas and Seanna M Martin contributed equally to this work.
Acute axonal degeneration
Central nervous system
Dorsal root ganglion
Enhanced green fluorescent protein
Glial cell line-derived growth factor
Posterior lateral line
Peripheral nervous system
Terminal deoxynucleotidyl transferase-mediated dUTP nick end labeling
We are grateful to Catalina Lafourcade and Florencio Espinoza for technical and administrative help, respectively. The following colleagues kindly provided fish strains or reagents: Herwig Baier, Hernán López-Schier, Darren Gilmour, Alex Nechiporuk, Phil Crosier, Graham Lieschke, Jau-Nian Chen. RV was funded by fellowships from CONICYT 21060426, 24080075 and 23110054; a traveling fellowship from MECESUP, the Company of Biologists, IBRO, Boehringher Ingelheim Fonds and the Vicerrectoría de Asuntos Académicos, Universidad de Chile. AS was funded by a grant from the National Institute of Dental and Craniofacial Research (NIDCR), KCO was supported by the UCLA Training Program in Neural Repair (funded by the National Institute of Neurological Disorders; NINDS 5/T32/NS07449:13), and SM was supported by a predoctoral fellowship from the NIDCR. MA was funded by FONDAP project number 15090007 and FONDECYT project number 1110275.
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