Postembryonic neuronal addition in Zebrafish dorsal root ganglia is regulated by Notch signaling
© McGraw et al.; licensee BioMed Central Ltd. 2012
Received: 14 April 2012
Accepted: 11 May 2012
Published: 27 June 2012
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© McGraw et al.; licensee BioMed Central Ltd. 2012
Received: 14 April 2012
Accepted: 11 May 2012
Published: 27 June 2012
The sensory neurons and glia of the dorsal root ganglia (DRG) arise from neural crest cells in the developing vertebrate embryo. In mouse and chick, DRG formation is completed during embryogenesis. In contrast, zebrafish continue to add neurons and glia to the DRG into adulthood, long after neural crest migration is complete. The molecular and cellular regulation of late DRG growth in the zebrafish remains to be characterized.
In the present study, we use transgenic zebrafish lines to examine neuronal addition during postembryonic DRG growth. Neuronal addition is continuous over the period of larval development. Fate-mapping experiments support the hypothesis that new neurons are added from a population of resident, neural crest-derived progenitor cells. Conditional inhibition of Notch signaling was used to assess the role of this signaling pathway in neuronal addition. An increase in the number of DRG neurons is seen when Notch signaling is inhibited during both early and late larval development.
Postembryonic growth of the zebrafish DRG comes about, in part, by addition of new neurons from a resident progenitor population, a process regulated by Notch signaling.
The neural crest, a transient population of vertebrate-specific progenitor cells, forms many disparate types of tissue after migration throughout the embryo (reviewed in [1, 2]). In the trunk, neural crest gives rise to the pigment cells of the skin and the neurons and glia of the peripheral nervous system (PNS), including the sensory neurons and glia of the dorsal root ganglia (DRG). The DRG are located in register with somites along the lateral edges of the spinal cord, and convey sensory information to the dorsal horn. The DRG are established from a population of neural crest cells that migrate along a medial pathway between somites and neural tube, to form discrete ganglia.
Mammalian and avian DRG contain three types of sensory neurons: those that relay information about touch and limb position (mechanoreceptive and proprioceptive neurons) and those that convey information about painful and irritating stimuli (nociceptive neurons). In mammals and birds, development of the DRG occurs in successive waves (reviewed in [3–5]), regulated through the activity of two neurogenin genes . The first wave depends upon the activity of Neurogenin2 (neurog2), and gives rise to large-diameter mechanoreceptive and proprioceptive neurons. The second wave of neurogenesis is Neurogenin1 (neurog1)-dependent and gives rise to mechanoreceptive and nociceptive neurons. A third wave of neurogenesis arises from migration and differentiation of Krox20-expressing boundary cap cells that differentiate into nociceptive neurons [7, 8]. After neurogenesis is complete, cell death regulated by soluble neurotrophins refines the neuronal cell population to reach its final size before the end of embryogenesis.
Sensory neurogenesis in the zebrafish occurs in a somewhat different manner. Differentiation of neurons in the DRG is dependent on the activity of only one Neurogenin gene, neurog1, and no equivalent of neurog2 exists in the zebrafish genome. DRG initially form with only two to five neurons differentiating from the neural crest by the end of embryogenesis. Adult animals have in the range of 100 neurons [10, 11], and thus the vast majority of neurons must be added after embryogenesis. Zebrafish trigeminal sensory neurons born at different times generate different classes of neurons , suggesting that the postembryonic growth of the DRG might reflect a similar increase in complexity.
Here, we describe a mechanism by which zebrafish DRG neurons are added after embryogenesis. We find that neurons are added steadily throughout larval development in a process regulated by Notch signaling. In mammals, a subset of neural crest stem cells reside in an undifferentiated state in many adult tissues, including the DRG (reviewed in ), leading us to hypothesize that similar cells might be the source of additional neurons in zebrafish. Using in vivo lineage tracing, we find that some new neurons are generated from progenitor cells that are resident in the DRG.
Proliferation of cells associated with the dorsal root ganglia (DRG)
Average number of cells per DRG at five dpf
BrdU incorporation at two dpf
BrdU incorporation at five dpf
3.26 ± 0.27
2.11 ± 0.47
0.09 ± 0.05 (4/144)
3.74 ± 0.20
1.98 ± 0.33
0.04 ± 0.02 (2/204)
3.86 ± 0.17a
2.67 ± 0.47
0.19 ± 0.05 (9/187)
4.32 ± 0.15b
1.69 ± 0.41
0.04 ± 0.04 (2/226)
Fates of labeled Tg(sox10:nlsEos) cells
To determine if proliferation is required for the increase in DRG neurons following Notch inhibition, we used acute or pulse-chase BrdU incorporation following the strategy described above. For the pulse-chase paradigm, larvae were treated with BrdU at two dpf and then incubated in DMSO or DAPT from two to five dpf. For acute BrdU experiments, larvae were exposed to DMSO or DAPT from two to five dpf and then treated with BrdU at five dpf just prior to fixation. Both DAPT treatment conditions showed a significant increase in the number of Elavl + neurons as compared to DMSO controls (Table 1). Following DAPT treatment, no significant increase was seen in BrdU + cells or BrdU+/Elavl + cells in either condition when compared to DMSO controls. These results suggest that blocking Notch signaling results in addition of DRG neurons primarily by promoting differentiation rather than increasing proliferation of progenitor cells.
Zebrafish, like other aquatic vertebrates, undergo substantial growth after embryogenesis is complete, including in the size and complexity of their nervous system. We demonstrate the addition of new neurons to the zebrafish DRG is continuous during postembryonic (larval) growth. Similar increases in DRG neuron number have been reported during amphibian metamorphosis . By contrast, growth of DRG in avian and mammalian species occurs during embryogenesis in overlapping waves that produce neurons of distinct function [6, 28–32]. It remains to be determined whether distinct neuronal classes are added to the zebrafish DRG at specific developmental stages. While postembryonic growth of DRG in mammals has been controversial, addition of neurons in other mammalian PNS structures has been well documented [33, 34].
Our lineage tracing experiments demonstrate that progenitors that reside among the Sox10-positive non-neuronal cells of the DRG are one source of new neurons during postembryonic growth. These results are consistent with a recent study demonstrating the addition of new DRG neurons from peripherally located precursors during avian embryogenesis . In zebrafish, this process appears to continue long after embryogenesis has ended. We demonstrate that zebrafish progenitor cells are associated with the zebrafish DRG after neural crest migration is complete, although these cells share Sox10 expression with the neural crest cells from which they are derived. Previous studies in mammals have suggested continued addition of neurons during late embryonic development from other sources, including neural crest-derived boundary cap cells found at the dorsal and ventral roots [35–37]. Boundary cap cells can form a number of late-developing sensory neurons [7, 8] and also acquire glial characteristics [38, 39]. We believe that the progenitors we have identified in zebrafish are distinct from boundary cap cells, as they are located in positions distinct from the dorsal and ventral roots, and do not express Krox20/egr2 mRNA (data not shown).
While sensory neurons are generally thought to be postmitotic, two previous studies have suggested that zebrafish DRG Elav + cells divide to give rise to new neurons [10, 18]. These results are in contrast to our findings and to those in other systems . We performed careful three-dimensional analysis to distinguish labeled cells, and note that while we have identified a very small number of Elav + cells that have incorporated BrdU, we do not find cells that are labeled with pH3. Furthermore, by time-lapse analysis we find that differentiated neurons do not divide, but rather arise de novo. We conclude that dividing Elav + cells are not a major source of DRG neurons in zebrafish.
We have not resolved whether DRG sensory neuron progenitors are bona fide glial cells, or whether they are a distinct population. Nervous system stem cells are now well established to have glial cell characteristics (reviewed in [40, 41]), including those found in the PNS. We identified potential DRG progenitors by continued expression of the Tg(sox10:nlsEos) transgene and their positions adjacent to DRG neurons. Whether these cells acquire characteristics of satellite glia, which surround sensory neuron cell bodies and alter neuronal function [42, 43], will require further study.
Our data suggests that Notch signaling regulates sensory neuron addition. We found that non-neuronal cells associated with differentiated DRG neurons express the Notch ligands deltaA and deltaD, as well as the Notch receptor notch1a. Inhibition of Notch signaling induces differentiation of DRG neurons in early- and late-stage larval fish. Our results are consistent with previous reports of an increase in DRG neuron number in zebrafish notch1a mutants . Recent reports have also demonstrated that loss of Notch signaling increases the initial differentiation of DRG neurons from neural crest [45, 46]. Our results suggest that Notch signaling normally prevents differentiation of neuronal progenitors. The expression of both ligand and receptor in subsets of cells adjacent to DRG neurons suggests a model where progenitor cells may act as an equivalence group: as cells begin to differentiate they inhibit their neighbors from doing so. One test of equivalence, whether cells are replaced by their neighbors after genetic or mechanical ablation, awaits future study. The methods we have used to block Notch signaling will act ubiquitously throughout the embryo, leaving open the possibility that effects are nonautonomous. In this scenario, blocking Notch would affect another cell type that would subsequently influence DRG development. Further experiments are needed to determine if Notch signaling acts cell autonomously within DRG progenitors.
Multiple roles for Notch signaling in neural crest development and differentiation have been previously identified, suggesting that this signaling pathway functions in a context-dependent manner. Notch signaling plays a role in the establishment of neural crest [17, 22, 47], and in the development of neural crest-derived cartilage and heart tissue . Multiple roles for Notch signaling have also been described for glial cell specification and differentiation [49–52]. Here we describe a later role for Notch in regulating the differentiation of neuronal progenitors during postembryonic phases of zebrafish DRG development. Similar roles have been previously described for Notch signaling in avian sensory and sympathetic progenitors  and mouse enteric progenitors . While the different roles for Notch signaling might be due to context or timing, the relative level of pathway stimulation may also have an effect . In addition, oscillating expression of Notch signaling components alters the effects of this signaling pathway in the embryonic brain . In cultures of neural crest stem cells, reduced levels of Notch signaling inhibit neurogenesis but do not promote gliogenesis, thus promoting self-renewal and maintaining pluripotency .
The presence of latent progenitors for postembryonic DRG growth is intriguing, and suggests a possible source of cells for regeneration. In mammals, neural crest-derived resident stem cells have been documented in a number of adult tissues, including amongst the enteric nervous system, heart, peripheral nerves and skin [58–64]. Neural crest stem or progenitor cells have also been isolated from embryonic and adult mammalian DRG [61, 65–70]. These cells may be involved in replacement or repair instead of growth, the possible evolutionary origin of their function.
Our work demonstrates the orderly addition of sensory neurons to the zebrafish DRG in the weeks after embryogenesis is complete, long after neural crest migration is complete. Neurons arise from dividing latent progenitor cells associated with the DRG that are amongst the satellite glial population. Progenitor differentiation is regulated by Notch signaling, throughout the period of larval development. The work provides insight into sensory neurogenesis in animals that undergo dramatic postembryonic growth.
Larval and adult zebrafish were maintained at 28.5°C in a 14 hour/10 hour light–dark cycle with twice daily feedings. Embryos were generated from natural crosses between adults and staged in hours post fertilization (hpf) as described by . Larval fish were staged in days postfertilization (dpf) and standard length . The transgenic lines used were: Tg(neurog1:EGFP) w61 described in ; TgBAC(neurod:EGFP) nl1 described in , Tg(sox10:nlsEos) w18 described in  and Tg(hsp70:XdnSu(H)myc) vu21 described in . Construction of the Tg(neurod:TagRFP) w69 transgenic line is described below. All work was approved by the University of Washington Institutional Animal Use and Care Committee.
The Tg(neurod:TagRFP) w69 transgenic line was constructed using the Multisite Gateway system . The p5E-neuroD construct was generated using a 5 kb region of the neurod 5′ promoter (; a gift from Teresa Nicolson). The p5E-neuroD construct was recombined with pME-TagRFP (a gift from Chi-Bin Chien) and p3E polyA. The construct was injected into one-cell embryos with Tol2 polymerase . Injected embryos were raised to sexual maturity and screened by pair-wise crosses to obtain germ line transgenic fish that generated the expected expression pattern. The final construct was injected into one-cell zygotes, which were then raised to maturity and screened for progeny that carried the Tg(neurod:TagRFP) transgene in DRG neurons amongst other cells.
For immunohistochemistry, larvae were collected at the stages indicated, euthanized in MS-222 (Sigma-Aldrich, St Louis, MO, USA); 10 mg/ml in buffered embryo medium) and fixed in 4% paraformaldehyde (PFA) in phosphate buffered saline (PBS) for two hours at room temperature. Antibody labeling was carried out as previously described . In brief, embryos were washed in PBS with 0.1% TritonX-100 (PBT), and blocked with the addition of 2% goat serum. Prior to blocking, fish were permeablized with three one-hour water washes. Fish were incubated in primary antibodies diluted in blocking solution overnight at room temperature (RT). Primary antibodies used were anti-GFP (1:700; rabbit or mouse anti-GFP; Invitrogen, Carlsbad, CA, USA), anti-Elavl (1:700; monoclonal antibody (mAB) 16A11; also called anti-HuC/D; ; Invitrogen); anti-myc tag mAB (1:500, Cell Signaling Technology, Beverly, MA, USA), anti-5-bromo-2-deoxyuridine (rat anti-BrdU; 1:100; Abcam, Cambridge, UK) and anti-phosphohistone H3 (rabbit anti-pH3; 1:100, Cell Signaling Technology). Fish were incubated in Alexa-488, Alexa-568 or Alexa-647 conjugated secondary antibodies (Invitrogen) overnight at room temperature, rinsed in PBT and then stored in 50% glycerol/PBS for imaging. Nuclei were visualized by DAPI labeling (Invitrogen).
Fluorescent RNA in situ hybridization was performed as described . Fixed larvae were made permeable for 20 minutes at RT using 10 μg/ml Proteinase K (Sigma-Aldrich) in PBT and then refixed for 15 minutes in 4% PFA. All hybridizations were carried out at 55°C. Digoxygenin-labeled antisense probes were generated using the following restriction enzyme and polymerases: XbaI/T7 for notch1a; EcoR1/T7 for deltaA and deltaD[79, 80]. Following in situ hybridization, larvae were processed for immunohistochemistry as described above. Larvae were then stored in 50% glycerol/PBS for imaging.
The pharmacological inhibitor of gamma-secretase activity N-N-(3,5-difluorophenacetyl)-L-alanyl]-S-phenylglycine t-butyl ester (DAPT; ; Sigma-Aldrich) was used to conditionally inhibit Notch signaling. DAPT was dissolved in dimethylsulfoxide (DMSO) at 10 mM and a working stock was diluted to 100 μM in embryo medium (EM) as previously described . Control fish were mock-treated with 1% DMSO in EM, for the same time course as the DAPT-treated fish. Zebrafish larvae were examined for the role of Notch signaling at two time points, either during early larval development, between two and five dpf, or during late larval development, between twenty and twenty-five dpf. Fish were treated continuously with DAPT or DMSO and were fed daily for the duration of the treatment. Following fixation, fish were processed for immunohistochemistry.
To corroborate the results seen with DAPT treatment, a heat-shock inducible dominant negative Suppressor of Hairless transgenic, Tg(hsp70l:XdnSu(H)myc), was used . Larvae were heat-shocked at 40°C for one hour twice daily between two and five dpf, fixed in 4% PFA, and processed for immunohistochemistry. Activation of the transgene was confirmed by the presence of myc labeling with an anti-myc antibody (Invitrogen).
To assess cell cycle in the DRG, we used 5-bromo-2-deoxyuridine (BrdU) incorporation to label cells in S-phase and anti-phosphohistone H3 (pH3) antibody to label cells in M-phase . In larvae, BrdU incorporation was carried out using a modified established protocol  Larvae were incubated in 10 mM BrdU dissolved in EM and 10% DMSO for 30 minutes at 4°C followed by one hour at 28.5°C. For pulse-chase experiments, larvae were exposed to BrdU between 50 and 51.5 hpf, transferred to DAPT or DMSO until five dpf and then fixed and processed. For pulse experiments, larvae that had been treated with DAPT or DMSO between two and five dpf, were exposed to BrdU at five dpf using the above protocol and then fixed and processed for BrdU detection. For BrdU incorporation in older larval fish, animals at 25 dpf were anesthetized in MS-222 and injected intraperitoneally using a glass capillary needle with 10 nl of a 10 mM BrdU in buffered Hank’s solution (protocol modified from ), incubated in EM at 28.5°C for two hours and then fixed for processing. BrdU detection was carried out using a previously describe protocol  Following BrdU detection, fish were immunolabeled with anti-Elavl and anti-pH3 antibodies and stored in 50% glycerol/PBS for imaging.
Homozygous larvae carrying the Tg(sox10:nlsEos) transgene were raised to four dpf, anesthetized with MS-222, and mounted in 0.8% agarose on a glass-bottomed cell culture dish. Cells surrounding the DRG were exposed to 405 nm light on an Olympus FV1000 confocal microscope (Olympus, Tokyo, Japan) to convert individual green nuclei to red. Following photoconversion, larvae were released from the agarose, and incubated in either 100 uM DAPT or DMSO as a control. At six dpf, larvae were fixed in 4% paraformaldehyde for two hours at room temperature and labeled with anti-ElavI antibody, mounted in Vectashield (Vector Laboratories, Burlingame, CA, USA) and imaged.
Prior to imaging, fish were mounted on bridged cover slips in 50% glycerol/PBS. Images and cell counts were obtained using a Zeiss LSM-5.0 Pascal confocal microscope (Carl Zeiss AG., Oberkochen, Germany), a Zeiss Axioplan 2 microscope and a Spot CCD camera (Diagnostic Instruments, Palo Alto, CA, USA), an Olympus FV1000 confocal microscope, or a 3I Marianas spinning disk microscope (Intelligent Imaging Innovations, Inc., Denver, CO, USA). Whole images were processed using ImageJ software (rsbweb.nih.gov/ij/index.html) and adjusted for brightness and contrast using Adobe Photoshop. For most individual cell counts, the five rostral-most DRG were analyzed in each condition. Non-neuronal cells were defined as associated with the DRG if they were in direct contact with labeled DRG neuronal cell bodies. Statistical significance was determined using Microsoft Excel software (Microsoft, Seattle, WA, USA) or GraphPad Prism version 5.0d software (Graphpad Software, San Diego, CA, USA).
For live imaging of neural crest migration and initial differentiation of neurons, embryos were anesthetized in MS-222, then embedded in 1.2% low melt agarose (Sigma-Aldrich) in EM. Cells were imaged every 10 minutes with a 20x water lens. In some cases stacks were assembled using Slidebook. Cells were tracked manually using ImageJ software. To image neuronal addition in larval fish, animal were anesthetized and embedded in 1.5% low-melt agarose and imaged using a 40x dipping lens. Following imaging, fish were released from the agarose and revived. Images were collected daily over a four-day time course.
5-difluorophenacetyl)-L-alanyl]-S-phenylglycine t-butyl ester
Dorsal root ganglion
Enhanced green fluorescent protein
Green fluorescent protein
Phosphate buffered saline
Tag red fluorescent protein.
We thank Bruce Appel for the Tg(hsp70:XdnSu(H)myc) line and for the sox10 promoter, Teresa Nicolson for the p5E-neuroD construct, and Chi-Bin Chien for the pME-TagRFP construct. We thank David White and fish facility staff for excellent fish care. Work was supported by National Institutes of Health grants R01 NS057220 (DWR), T32 GM007270 (AP) and T32 HD007183 (HFM).
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