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. 2011 Jan 16;218(3):271–276. doi: 10.1111/j.1469-7580.2010.01337.x

Somatosensory mechanisms in zebrafish lacking dorsal root ganglia

Yasuko Honjo 1, Laurel Payne 1, Judith S Eisen 1
PMCID: PMC3058213  NIHMSID: NIHMS280431  PMID: 21235542

Abstract

Dorsal root ganglion (DRG) sensory neurons transmit all somatosensory information from the trunk region of the body. erbb3 mutant zebrafish do not form DRG neurons because the neural crest cells that generate them migrate aberrantly. Here we report that homozygous erbb3 mutants appear to swim and feed normally, and that they survive through adulthood, despite never forming DRG neurons. The source of sensory compensation in adult erbb3 mutants remains unknown, although it may be from lateral line ganglion neuromasts which are reduced, but present, in erbb3 mutants. We also provide new information about the development of DRG neurons in wild-type juvenile zebrafish.

Keywords: DRG, erbb3, somatosensory, zebrafish

Introduction

Somatosensory information such as touch, pain, temperature, and proprioception is vital for survival. For example, the sensation of pain promotes escape from the source of pain, such as predators. In the trunk region of vertebrates, DRG neurons are responsible for transmitting somatosensory information to the central nervous system (CNS) (Scott, 1992; Lewin & Moshourab, 2004). DRG neurons are generated by neural crest cells (Le Douarin & Kalcheim, 1999). In zebrafish, a subset of migrating neural crest cells stall adjacent to the neural tube and later these cells generate DRG neurons and glia (Raible et al. 1992). Another neural crest derivative, boundary cap cells, has also been reported to contribute cells that later generate some DRG neurons in mice (Golding & Cohen, 1997; Maro et al. 2004; Hjerling-Leffler et al. 2005). DRGs consist of several classes of neurons that are functionally and morphologically distinct, as well as glial cells (Scott, 1992). DRG neurons are an excellent model system in which to study neural development, including specification of distinct cell fates and elaboration of appropriate cell-type specific properties. DRGs are also an excellent model in which to learn how sensory information is received and conveyed to the CNS.

In addition to DRG neurons, anamniote vertebrates, such as zebrafish, have an earlier developing population of cells, Rohon-Beard (RB) neurons, that provide sensory information in the trunk (Reyes et al. 2004). RB neurons reside in the spinal cord and are generated from the same cell population that gives rise to neural crest cells (Cornell & Eisen, 2000). In zebrafish, as in most anamniote species studied to date, the entire population of RB neurons dies and is functional replaced by DRG neurons. Interestingly, lack of DRG neurons does not lead to RB survival, suggesting that the two cell populations develop independently (Reyes et al. 2004). As zebrafish DRGs develop, they are typically first populated by a single neuron (Raible et al. 1992; An et al. 2002); the number of neurons increases over the time (An et al. 2002). However, details about specification of DRG neuronal subclasses are lacking for zebrafish.

The lateral line is another sensory system in fish and amphibians (Ghysen & Dambly-Chaudiere, 2007). Lateral line neurons are located along the surface of the mid-region of the body. This sensory system transmits information about changes in water current (Ghysen & Dambly-Chaudiere, 2007). The lateral line sensory system works together with DRGs and other sensory systems to obtain and transmit environmental information that leads to adjustment in body movement. Lateral line axons migrate from the cranial region to their target neuromast hair cells in the skin. Migrating lateral line axons co-migrate with glial cells (Gilmour et al. 2002).

Given the importance of sensory innervation, we were surprised to find that zebrafish erbb3 mutants, which lack DRGs (Honjo et al. 2008) and have defects in pigmentation (Budi et al. 2008) and lateral line glia (Lyons et al. 2005), survive through adulthood and show at least some normal behaviors, for example, they appear to swim normally and are able to pursue food in the water column. ErbB3 is a receptor tyrosine kinase of the EGF receptor family and is involved in a variety of biological processes including glial migration and development (Wells, 1999; Olayioye et al. 2000; Corfas et al. 2004; Holbro & Hynes, 2004). Here we describe our studies of zebrafish erbb3 mutants to learn how they can survive and show at least some normal behaviors in the absence of DRG neurons. This study expands our knowledge of development of DRG and other sensory neurons in juvenile and adult zebrafish, and addresses possible sources of compensation for the missing DRG somatosensory system.

Materials and methods

Animals

Embryos were obtained from natural spawnings of a wild-type (AB) colony or crosses of identified carriers heterozygous for specific mutations; the mutations used were erbb3bst48 (Lyons et al. 2005) and erbb3utr4e1 (Budi et al. 2008). We also used the –3.1ngn1:Green Fluorescent Protein (GFP) transgenic line (Blader et al. 2003), which we refer to as Tg(ngn1:GFP). Fish were maintained in the University of Oregon Zebrafish Facility on a 14 h light/10 h dark cycle at 28.5 °C. Embryos and larvae were staged according to Kimmel et al. (1995) by days post fertilization at 28.5 °C (dpf), or weeks post fertilization (wpf) at 28.5 °C as well as by body length (BL, measured from snout to tip of tail). We also estimated standardized standard length (eSSL) according to Parichy et al. (2009). Body length was well correlated with age and not significantly different between wild types and erbb3 mutants at 7 dpf (wt: 3.0–4.0 mm, average 3.8 mm; erbb3 mutants: 3.2–4.0 mm, average 3.8 mm; e3.5 SSL), 2 wpf (wt: 6.5–8.0 mm, average 7.1 mm; erbb3 mutants: 6.5–7.5 mm, average 7.2 mm; e6.2–e7.2 SSL), 4 wpf (wt: 16.5–18.0 mm, average 17.6 mm; 15.0–19.5 mm, average 17.6 mm; e10.4–e11.0 SSL) or 6 wpf (wt: 26.0–29.0 mm, average 27.1 mm; erbb3 mutants: 24.0–29.0 mm, average 26.6 mm; e23 SSL), although erbb3 mutants had a slightly lower BL at 8 wpf (wt: 29.0–31.0 mm, average 29.8 mm, e25 SSL; erbb3 mutants: 27.0–31.0 mm, average 28.1 mm, e24 SSL) and 10 wpf (wt: 30.0–32.0 mm, average 30.5 mm, e26 SSL; erbb3 mutants: 28.0–30.0 mm, average 28.6 mm, e25 SSL).

Antibodies

Anti-Elavl antibody (16A11) was used at a ratio of 1 : 1000 (Marusich et al. 1994; Henion et al. 1996) and anti-acetylated tubulin antibody (Sigma) at 1 : 200 (Ungos et al. 2003). The zn5 and zn12 antibodies were used at a ratio of 1 : 1000 and 1 : 250, respectively (Fashena & Westerfield, 1999; Menelaou et al. 2008). Anti-trkB (Santa Cruz Inc.) was used at a ratio of 1 : 100 (Germana et al. 2004). Alexa-488 conjugated goat anti-mouse monoclonal antibody was used as the secondary antibody. Anti-S100 (Dako) was used at a ratio of 1 : 1000 (Germana et al. 2008).

Cryosection immunohistochemistry

Animals were anesthetized in 0.01% MS222 (3-aminobenzoic acid ethyl ester), fixed either intact or in pieces with 4% paraformaldehyde (PFA) for 2 days at 4 °C, and cryosectioned. Sections were incubated with phosphate-buffered saline (PBS) followed by blocking solution [PBS, 1% dimethylsulfoxide (DMSO) and 0.1% Triton and 2% goat serum), then incubated with primary antibody in blocking solution overnight at 4 °C. After washing with PBS, sections were incubated with secondary antibody in blocking solution for 5 h at room temperature (RT) or overnight at 4 °C, washed with PBS, and mounted for observation.

Slice immunohistochemistry

For slice staining, fish were fixed with 4% PFA overnight and sliced into three pieces with a razor blade. Each slice was fixed again overnight. Immunohistochemistry for slices was performed as in Ungos et al. (2003) and Olsson et al. (2008). Briefly, animals were anesthetized in 0.01% MS222 fixed in 4% formaldehyde for 3 h at RT, then washed in PBS. Fixed whole embryos and larvae were permeabilized in distilled water for 3 h before being incubated with blocking solution for 1 h. Preparations were incubated with primary antibody overnight at RT, washed with PBS, then incubated with secondary antibody overnight at RT and washed with PBS.

Lateral line staining

Neuromast hair cells were labeled by incubating live larvae or adults in 125 μm 4-(4-diethylaminostyryl)-N-methylpyridinium iodide (4-Di-2-Asp) in fish Ringer's solution for about 3 h (Alexandre & Ghysen, 1999).

Results

To begin to describe subtypes of DRG neurons, we cryosectioned fish every other week from 2 wpf through 10 wpf and stained them with a battery of antibodies, including anti-Elavl, anti-acetylated tubulin, anti-TrkB (Fig. 1), anti-S100, and zn5 (data not shown). We found morphologically and immunohistologically distinct subtypes of DRG neurons located adjacent to the spinal cord in wild-type fish (Fig. 1). For example, only a subpopulation of DRG neurons expressed TrkB, as described in other species (Rifkin et al. 2000; Kramer et al. 2006). We also found the number of cells in the DRG increased over time in wild types, as previously described (Fig. 1; An et al. 2002).

Fig. 1.

Fig. 1

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erbb3 mutants lack DRG neurons. (A,C,E,G) Images of anti-Elavl antibody staining, (I) image of anti-acetylated tubulin (AcTub) antibody staining, (K) image of anti-TrkB antibody staining and (B,D,F,H,J,L) overlays of fluorescent and differential interference contrast (DIC) images. The number of GFP-positive DRG neurons (arrowheads) increased over time in 2-wpf (A,B), 4-wpf (C,D) and 10-wpf (E,F) wild-type fish. In contrast, GFP-positive DRG neurons were not present in 6-wpf erbb3 mutants (G, H). By 10 wpf, wild-type DRG neurons have differentiated into morphologically and immunohistologically distinct subpopulations, as shown by anti-acetylated tubulin (I,J) and anti-TrkB antibodies (K,L). Scale bar: 100 μm (A,B in B; C–L in D).

The number of DRG neurons normally increases over time in zebrafish (Raible et al. 1992; An et al. 2002); however, the precise source of the new cells has not been described. Therefore, we postulated that erbb3 mutants might initially lack DRG neurons, but that some unknown source of cells could enable DRG neurons to form later in development. To test this possibility, we examined erbb3 mutants between 2 and 10 wpf using the same battery of markers we used on wild types. Contrary to our hypothesis, none of these markers labeled cells in the position of DRGs in erbb3 mutants (Fig. 1). These results suggest that DRG neurons do not form in erbb3 mutant zebrafish. These results also suggest that the later generated DRG neurons in wild-type zebrafish do not arise from a cryptic cell population, but likely arise from proliferation of neural crest-derived cells located at the site where DRGs develop.

Because erbb3 mutants appear never to form DRGs, we next examined whether RB neurons might persist in these mutants and thus compensate for lack of DRG neurons. We used the Tg(ngn1:GFP) line that was previously reported to express GFP in RBs (Blader et al. 2003). We found that at 7 dpf, wild-type fish still had a few RB neurons; however, these cells had all died by 4 wpf (Fig. 2). Similarly, RB neurons were also present in 7 dpf erbb3 mutants, but these cells were entirely gone by 4 wpf. We also found that RBs were absent from cryosectioned erbb3 mutants labeled with anti-Elavl, zn5 or zn12 antibodies (data not shown). These results provide evidence that RB neurons die in erbb3 mutants as they do in wild types, and thus that they are not available to compensate for the loss of DRG sensory mechanisms.

Fig. 2.

Fig. 2

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Rohon-Beard neurons die normally in erbb3 mutants. (A,C,E,G) images of anti-GFP antibody staining and (B,D,F,H) overlays of fluorescent and DIC images. GFP-positive RB neurons (arrowheads) were present in 1-wpf Tg(ngn1:GFP) fish (A, B) as well as Tg(ngn1:GFP) fish crossed with erbb3 mutants (C, D). However, by 4 wpf, both wild-type Tg(ngn1:GFP) fish (E,F) and erbb3 mutant fish crossed with Tg(ngn1:GFP) fish (G,H) lack RB neurons suggesting that RB neurons die normally in erbb3 mutants. Scale bar 100 μm (A–D in B) and 120 μm (E–H in F).

DRG neurons extend axons toward a variety of targets, including skin and fin (Reyes et al. 2004). We hypothesized that by examining whether axonal processes were present in these locations in erbb3 mutants, we might be able to learn the source of cells that compensate for DRG functions. To learn whether any axonal processes innervate the skin or the fin in erbb3 mutants, we performed anti-acetylated tubulin antibody staining on 2 and 4 wpf fish. We found that acetylated tubulin-positive processes were present in the skin on the inner surface of the scales of both wild types and erbb3 mutants (Fig. 3). To learn whether these processes were from DRG axons, we examined Tg(ngn1:GFP) fish. GFP-positive axonal processes were present in the skin of wild-type fish and appear to be similar (Fig. 3). However, there were no GFP-positive axonal processes in erbb3 mutants (data not shown). These results suggest that the processes in the skin of wild types emanate from DRG neurons, and are consistent with our finding that DRG neurons never form in erbb3 mutants.

Fig. 3.

Fig. 3

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Acetylated tubulin (AcTub)-positive neuronal processes on the skin are similar to the GFP-positive neuronal processes on the skin in Tg(ngn1:GFP) fish. (A) Anti-acetylated tubulin staining in 4 wfp wild-type fish. The neuronal processes look the same as the GFP-positive neuronal processes in Tg(ngn1:GFP) fish (B). Similar Acetylated tubulin-positive neuronal processes are present in erbb3 mutants (C). Scale bar 120 μm.

Because DRG neuron processes were absent from erbb3 mutants, we next examined whether any of the processes in the skin emanated from lateral line nerves. Lateral line glia migrate aberrantly along lateral line nerves in erbb3 mutants (Lyons et al. 2005). Despite the lack of glial cells, lateral line nerves were still present in 4 wpf erbb3 mutants; however, projections to neuromasts were defasciculated, as previously described (Lyons et al. 2005). We examined whether the hair cells in erbb3 mutants were functionally normal by labeling them with 4-di-2-Asp. The number of neuromast hair cells was slightly reduced in erbb3 mutants; however, the cells were present (Fig. 4). Our finding that hair cells of erbb3 mutants were labeled with this dye suggests that they have a normal function (Alexandre & Ghysen, 1999; Gilmour et al. 2002; Ghysen & Dambly-Chaudiere, 2007). 4-di-2-Asp dye labeling also revealed lateral line afferent nerves innervating neuromast cells (Fig. 5). The morphology of lateral line nerve endings was a relatively symmetric extension toward neuromast cells with thicker central axon bundles. In contrast, the endings of the acetylated tubulin-positive neuronal processes on the skin were asymmetrically branched and all of the processes were very fine. Based on the differences in morphology of the nerve endings between lateral line neurons and the endings innervating the skin, the lateral line nerve endings did not appear to be the same as the nerve endings we observed with anti-acetylated tubulin on the inside surface of the skin in erbb3 mutants. This suggests that the lateral line sensory system is present and functioning in erbb3 mutants. However, the axonal processes we observed in erbb3 mutants are unlikely to be from lateral line nerves.

Fig. 4.

Fig. 4

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Neuromast hair cells are reduced, but present in erbb3 mutants. 4-di-2-Asp staining shows functional neuromasts in 4 wpf wild-type fish (A). The number of functional neuromasts is reduced in erbb3 mutants, but functional neuromasts remain present (B). Scale bar: 500 μm.

Fig. 5.

Fig. 5

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4-di-2-Asp staining reveals that the neuronal processes on the inner surface of scales in Tg(ngn1:GFP) fish are not from lateral line nerves innervating neuromasts. (A) Lateral line nerves innervating neuromasts (arrowhead). (B) Neuronal processes on the scales (asterisk) have a different morphology from lateral line nerves (arrowhead). Scale bar: 80 μm.

Discussion

Here we describe development of DRG neurons in juvenile zebrafish. We found that erbb3 mutants that lack DRG neurons as embryos and larvae do not form DRGs as juveniles. This suggests that the addition of DRG neurons in zebrafish larvae (An et al. 2002) is from neural crest-derived cells located at the position of the DRG, and that there is no additional generation of DRG neurons from a cryptic source in zebrafish. We also examined the development of lateral line nerves in juveniles. We found that in erbb3 mutants, at least some lateral line neurons survive without glial cells.

Compensation for DRG function

Surprisingly, erbb3 mutants survive through adulthood and show at least some behaviors that appear normal, such as swimming and feeding, without DRG neurons and in the absence of normal Schwann cell myelination (Lyons et al. 2005). To investigate possible sources of compensation for DRG neurons, we examined RB neurons and lateral line neuromasts. It is also possible that trunk DRG function is compensated by sensory information from cranial trigeminal sensory neurons, although we did not investigate this possibility.

Lateral line and processes on the skin

We analyzed the presence of innervation to skin using acetylated tubulin staining. We identified afferent lateral line nerves on the skin that innervate neuromasts. However, we also found some processes that were not afferent lateral line nerves, revealed by costaining with 4-di-2-Asp dye. We tried to label retrogradely from the skin using both DiI and rhodamine dextran to learn the source of these cells, but never detected any dye-labeled cell bodies. Thus, it is still unclear what type of neurons provides the fine arbors on the skin surface in erbb3 mutants. It is possible that there are axons from a very few remaining DRG neurons that we did not detect in cryosectioned samples. Similarly, the function of these processes is a question that needs to be addressed in the future. Lack of glial cells typically causes neuronal cell death (Jessen & Mirsky, 1999). However, although erbb3 mutants have defective migration of lateral line glia, lateral line nerves are present and appear functional. This may be because the cell bodies of afferent lateral line nerves remain in the head (Raible & Kruse, 2000; Ledent, 2002), where they may possibly be supported by glia that are unaffected by the absence of ErbB3.

Acknowledgments

We thank Will Talbot for erbb3b mutants, Uwe Strahle for the Tg(ngn1:GFP) line, Jacob Lewis and the University of Oregon Zebrafish Facility staff for animal husbandry, the University of Oregon Histology Facility for sectioning, Liesl Van Ryswyk and Christie Ojiaku for participation in section staining. Supported by NIH grants HD22486 and NS23915 and the Medical Research Foundation of Oregon.

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