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Keywords:

  • tyrosine kinase receptor;
  • substance P;
  • calcitonin gene-related peptide;
  • neurofilament;
  • phenotype;
  • neuron;
  • dorsal root ganglion

Abstract

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. LITERATURE CITED

The neuropeptide-immunoreactive (IR) and neurofilament-IR neurons are two major phenotypical classes in dorsal root ganglion (DRG). Tyrosine kinase receptor (Trk)A, TrkB, and TrkC are three members of the Trk family which may be relevant to neuronal phenotypes. Whether target skeletal muscle cells generate their expression remains unclear. Neurons containing substance P (SP), calcitonin gene-related peptide (CGRP), neurofilament 200 (NF-200), TrkA, TrkB, and TrkC were quantified using immunohistochemistry in rat DRG neuronal cultures and cocultures of DRG neurons and skeletal muscle cells. The percentage of NF-200 and TrkC-expressing neurons in cocultures of DRG neurons and skeletal muscle cells was significantly higher, 26.86% ± 3.17% (NF-200) and 27.74% ± 3.63% (TrkC) compared with 20.92% ± 1.98% (NF-200) and 16.70% ± 3.68% (TrkC) in DRG cultures; whereas the percentage of SP, CGRP, TrkA, and TrkB-expressing neurons was not changed significantly by the addition of target skeletal muscle cells. Thus, target skeletal muscle cells may influence neurofilament-phenotype and TrkC receptor but not neuropeptide-phenotype and TrkA and TrkB receptors. Anat Rec, 2009. © 2008 Wiley-Liss, Inc.

Targets of neuronal innervation play a vital role in regulating the survival and differentiation of innervating neurotrophin-responsive neurons (Howe et al.,2001). However, it is still largely unclear whether target skeletal muscle cells influence dorsal root ganglion (DRG) neuronal phenotyes and tyrosine kinase receptors (Trk) expression.

The neuropeptide-immunoreactive (IR) and neurofilament-IR neurons are two major phenotypical classes in DRG. Neuropeptide-IR neurons are considered to be with unmyelinated or thinly myelinated nociceptive afferents which are considered to innervate skin and viscera. Neurotransmitter expression in neurons is regulated by innervated target tissue and intrinsic neuronal properties (Braun et al.,1996). Cultured neurons expressed neuropeptides with a time course and in proportions similar to those in vivo (Hall et al.,1997). Neurofilament-IR neurons typically have myelinated axons which are considered to innervate muscle spindle. Neurofilament-IR neurons are also present in DRG cell cultures (Hall et al.,1997). It has been identified that DRG neurons express three Trk receptors (TrkA, TrkB, and TrkC) that only minimally overlap. TrkA-expressing neurons are mostly small cells with unmyelinated axons and are thought to innervate nociceptors and thermoreceptors. TrkC-expressing neurons are mostly large cells with myelinated axons and likely receive input from peripheral mechanoreceptors. TrkB-expressing cells vary from small to large and are generally medium sized and are likely to have myelinated axons (Wright and Snider,1995). The activation of Trk receptors seems to be essential in most cases for the biological effects of neurotrophins (Barbacid,1994).

Target tissues are essential for the maintenance of the function of neurons and nerve–muscle communication. In the absence of limb-derived trophic signals, the affected neurons fail to survive. Limb-bud targets removal results in the loss of spinal sensory neurons that normally innervate the limbs in the chick embryo. Sensory neurons lost in the absence of target-derived trophic support should be rescued by treatment with muscle extract or neurotrophic factors which are known to be required for the survival of these neurons (Snider,1994; Caldero et al.,1998). Extracellular application of skeletal myosin II to embryonic sensory neurons resulted in increasing axon formation and branching in each neuron in vitro (Silver and Gallo,2005). It has been shown that the increase in excitability of skeletal muscle is regulated by the excitatory afferents in the spinal cord-DRG-skeletal muscle organotypic coculture system (Streit et al.,1991).

There is some evidence for phenotypic plasticity in sensory neurons (Hall et al.,1997). Outgrowing sensory neurons appear to have more flexibility in their pathway and target choices than do motoneurons in vivo (Wang and Scott,1999). However, much less is known about the influence of target muscle cells on DRG neuronal phenotyes and Trk receptors. Here, we have established a neuromuscular coculture model of DRG neurons and skeletal muscle cells to test what extent to the expression of substance P (SP), calcitonin gene-related peptide (CGRP), neurofilament 200 (NF-200), TrkA, TrkB, and TrkC in DRG neurons in the presence of target skeletal muscle cells when compared with that in DRG culture in the absence of skeletal muscle cells.

MATERIALS AND METHODS

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. LITERATURE CITED

Cell Cultures

DRG cell culture preparations utilized embryonic 12.5-day-old Wistar rats maintained in the Experimental Animal Center at Shandong University of China. Under aseptic conditions, the bilateral DRG was removed from each embryo, placed in culture medium, and minced into fragments ∼0.5 mm in diameter, digested with 0.25% trypsin (Sigma) in D-Hanks solution at 37°C for 10 min, centrifuged, and triturated in growth media supplemented with 5% fetal bovine serum (Gibco). Dissociated DRG cells were plated at a density of 2×105 cells/mL in 6-well clusters (Costar, Corning, NY) which would contain 24 mm diameter coverslips precoated with poly-L-lysine (0.1 mg/mL) and then incubated at 37°C in a 5% CO2 incubator.

Skeletal muscle cell culture preparations utilized newborn Wistar rats maintained in the Experimental Animal Center at Shandong University of China. Skeletal muscle cell cultures were prepared separately from, but simultaneously with, the primary cultures of DRG cells. Under aseptic conditions and using the newborn rat, skeletal muscle was removed from the hind limb of each animal, minced with fine dissecting scissors into fragments ∼0.5 mm in diameter, digested with 0.25% trypsin (Sigma) in D-Hanks solution at 37°C for 20 min, centrifuged, and triturated in growth media supplemented with 5% fetal bovine serum (Gibco). Isolated skeletal muscle cells were plated at a density of 2×105 cells/mL in flasks (Costar, Corning, NY) and then incubated at 37°C in a 5% CO2 incubator for ∼2 hr to remove nonskeletal muscle cells.

At this time, the neuromuscular cocultures were prepared as follows. The skeletal muscle cell suspension was transferred from the flasks to the 6-well clusters which would contain dissociated DRG cells. Cultures of dissociated DRG cells alone were maintained continuously in culture media. All these culture preparations were incubated at 37°C in a 5% CO2 incubator for 6 days with media change every 2 days.

The composition of the culture media is DMEM/F-12 (1:1) supplemented with 10% fetal bovine serum, 2% B-27 supplement (Gibco), L-glutamine (0.1 mg/mL, Sigma), insulin (0.25 μg/mL, Sigma), penicillin (100 U/mL), and streptomycin (100 μg/mL).

Double Fluorescent Labeling of MAP2 and SP, CGRP, NF-200, TrkA, TrkB, or TrkC

At 6 days of culture age, DRG cultures and neuromuscular cocultures were processed for double immunofluorescent labeling of microtubule associated protein 2 (MAP2) and SP, CGRP, NF-200, TrkA, TrkB, or TrkC. The cells on coverslips were rinsed quickly one time in 0.1 mol/L Sorenson's phosphate buffer to remove media. The cells were fixed in 4% paraformaldehyde, pH 7.4, for 20 min at 4°C. After washing in 0.1 mol/L phosphate buffer saline (PBS) for three times, the cells were blocked by 2% normal goat serum in 0.1% Triton PBS to block nonspecific sites and permeates cells. The samples were incubated with rabbit polyclonal anti-SP, CGRP, NF-200, TrkA, TrkB, or TrkC (1:500) overnight at 4°C, respectively. After washing in 0.1 mol/L PBS three times, the samples were incubated by goat anti-rabbit conjugated to Cy3 (1:200) for 45 min in dark. After washing for three times in 0.1 mol/L PBS, the cells were incubated with mouse monoclonal anti-MAP2 (1:200) for 60 min in dark. After washing three times in 0.1 mol/L PBS, the cells were incubated with goat anti-mouse conjugated to Cy2 (1:200) for 45 min in dark. After washing in 0.1 mol/L PBS, the cells were coverslipped immediately with Vectashield anti-fade mounting media (Vecto Laboratories, Inc.) and stored at 4°C until observation by fluorescent microscope.

Quantitative Analysis of the Percentage of Neuropeptide-, NF-200-, and Trk-expressing Neurons

SP-IR, CGRP-IR, NF-200-IR, TrkA-IR, TrkB-IR, or TrkC-IR neurons were observed under a fluorescent microscope (Nikon) with 20× objective lens. SP-IR, CGRP-IR, NF-200-IR, TrkA-IR, TrkB-IR, or TrkC-IR neurons in five visual fields in the central part of each coverslip were counted as the positive neurons in each sample. The number of total neurons (MAP2-IR) were also counted in the same visual field. Then, the percentage of SP-IR, CGRP-IR, NF-200-IR, TrkA-IR, TrkB-IR, or TrkC-IR neurons would be obtained.

Statistical Analysis

Data are expressed as mean ± SD. Statistical analysis was evaluated with SPSS software by one-way ANOVA followed by the Student-Newman-Keuls test for significance to compare the differences among various groups. Significance was accepted at P < 0.05.

RESULTS

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. LITERATURE CITED

The Effects of Skeletal Muscle Cell on DRG Neuronal Phenotyes

To test the effects of skeletal muscle cells on SP, CGRP, and NF-200 expression in DRG neurons, DRG cells were cultured for 6 days with or without target skeletal muscle cells and processed for double fluorescent labeling of MAP2 and SP, CGRP, or NF-200, and then, DRG neurons containing SP, CGRP, and NF-200 were quantified. 117 SP-IR, 110 CGRP-IR, and 192 NF-200-IR neurons are in 914, 1121, and 910 total DRG neurons, respectively, in the absence of skeletal muscle cells. 130 SP-IR, 114 CGRP-IR, and 279 NF-200-IR neurons are in 947, 1066, and 1053 total DRG neurons, respectively, in the presence of skeletal muscle cells. The target skeletal muscle cells could promote NF-200 expression but not neuropeptides. 26.86% ± 3.17% of DRG neurons expressed NF-200 in neuromuscular cocultures which is higher than that in DRG cultures alone (20.92% ± 1.98% NF-200-expressing neurons of total cells). The percentage of SP-IR (13.63% ± 1.85%) and CGRP-IR (11.00% ± 3.93%) neurons in neuromuscular cocultures is not changed significantly as compared with that in DRG cultures alone (12.65% ± 2.38% SP-IR and 10.2% ± 5.87% CGRP-IR neurons) (Fig. 1).

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Figure 1. Panels A and B are double-fluorescent labeling of MAP2 and NF-200 in DRG neurons in absence or presence of skeletal muscle cells at 6 days of culture age (Scale bar = 50 μm). Panel A is DRG cultures in absence of skeletal muscle cells (A1: NF-200; A2: MAP2; A3: overlay of A1 and A2). Panel B is cocultures of DRG neurons and skeletal muscle cells (B1: NF-200; B2: MAP2; B3: overlay of B1 and B2). Panel C is the percentage of SP-, CGRP-, and NF-200-expressing DRG neurons in absence or presence of skeletal muscle cells at 6 days of culture age (n = 5). *P < 0.01.

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The Effects of Skeletal Muscle Cell on Trk Expression in DRG Neurons

To test the effects of skeletal muscle cells on Trk receptors expression in DRG neurons, DRG cells were cultured for 6 days with or without target skeletal muscle cells and processed for double fluorescent labeling of MAP2 and TrkA, TrkB, or TrkC, and then, TrkA-IR, TrkB-IR, or TrkC-IR DRG neurons were quantified. 232 TrkA-IR, 95 TrkB-IR, and 158 TrkC-IR neurons are in 910, 905, and 933 total DRG neurons, respectively, in the absence of skeletal muscle cells. 271 TrkA-IR, 111 TrkB-IR, and 269 TrkC-IR neurons are in 974, 949, and 962 total DRG neurons, respectively, in the presence of skeletal muscle cells. The target skeletal muscle cells could promote TrkC expression but not TrkA and TrkB. DRG neurons (27.74% ±3.63 %) expressed TrkC in neuromuscular cocultures which is higher than that in DRG cultures alone (16.7% ± 3.68% TrkC-expressing neurons of total cells). The percentage of TrkA-IR (27.75% ± 1.91%) and TrkB-IR (11.61% ± 1.88%) neurons in neuromuscular cocultures is not changed significantly when compared with that in DRG cultures alone (25.30% ± 3.00% TrkA-IR and 10.31% ± 3.09% TrkB-IR neurons) (Fig. 2).

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Figure 2. Panels A and B are double fluorescent labeling of MAP2 and TrkC in DRG neurons in absence or presence of skeletal muscle cells at 6 days of culture age (Scale bar = 50 μm). Panel A is DRG cultures in absence of skeletal muscle cells (A1: TrkC; A2: MAP2; A3: overlay of A1 and A2). Panel B is cocultures of DRG neurons and skeletal muscle cells (B1: TrkC; B2: MAP2; B3: overlay of B1 and B2). Panel C is the percentage of TrkA-, TrkB-, and TrkC-expressing DRG neurons in absence or presence of skeletal muscle cells at 6 days of culture age (n = 5). *P < 0.01.

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DISCUSSION

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. LITERATURE CITED

The aim of this study was to approach the question of neuronal phenotyes and Trk receptors expressing dependence on target skeletal muscle cells during development. As previous reports, multiple appropriate sensory neuron phenotypes arise in a regulated fashion in cultured neurons isolated before target connections have formed, and some candidate target tissues can modulate that intrinsic expression pattern (Hall et al.,1997). The identification of the cell types that express the Trk receptors is of particular interest because neurons subserving specific sensory modalities depend on distinct neurotrophins for survival during development (Stephens et al.,2005). In this study, we found that: (1) the percentage of DRG neurons contained neurofilament increased significantly in neuronmuscular cocultures of DRG neurons and target skeletal muscle cells as compared with that in DRG neuronal cultures, whereas the percentage of DRG neurons contained SP or CGRP in neuronmuscular cocultures was not changed significantly as compared with that in DRG cultures and (2) the percentage of TrkC-expressing DRG neurons increased significantly in neuronmuscular cocultures of DRG neurons and target skeletal muscle cells when compared with that in DRG neuronal cultures, whereas the percentage of TrkA- or TrkB-expressing DRG neurons in neuronmuscular cocultures was not changed significantly when compared with that in DRG cultures.

It has been shown that the developmental regulation of sensory neurons containing CGRP that predominantly contact visceral and cutaneous peripheral target end organs in vivo (Hall et al.,1997), whereas abundant neurofilament expression is characteristic of large neurons that innervate muscle spindles (Lawson et al.,1984; Perry et al.,1991). The results in this study provide an important demonstration that neurofilament-phenotype but not neuropeptide-phenotype could be regulated by the presence of target skeletal muscle cells. These results are consistent with that SP-IR and CGRP-IR neurons did not require an intact DRG or connections with other tissues to regulate neuropeptides often expressed by nociceptive neurons (Hall et al.,1997). In the present experiment, DRG was dissected out from embryonic rats on embryonic days 12.5 (E12.5) before neuropeptides appear in DRG neurons (Hall et al.,1997). At E12.5, many DRG neuron precursors are undergoing their final mitoses, and most have not extended peripheral processes (Lawson et al.,1984; Mirnics and Koerber,1995). These results implicated that some neurons in the embryonic DRG seem to be intrinsically specified to later express SP and CGRP. Neurofilament-IR neurons typically have myelinated axons which are considered to innervate muscle spindle (Hall et al.,1997). In DRG and skeletal muscle fibers cocultures, peripheral neurites of DRG neurons coiling around regenerated muscle fibers correspond to that part of the sensory muscle spindle apparatus which developed in vivo (Spenger et al.,1991). Our results are in agreement with the earlier previous reports. Neurofilament expression is dependent on, at least in part, the presence of target skeletal muscle cells.

Activation of receptor trkA seems important for the survival of cutaneous and visceral afferents, whereas trkC may be more important for muscle afferents (Hory-Lee et al.,1993; Crowley et al.,1994; Klein et al.,1994; McMahon et al.,1994; Smeyne et al.,1994; Hall et al.,1997). It has been suggested that the distribution of TrkC is necessary for the maintenance of the DRG proprioceptive neurons during embryonic development in mice (Stephens et al.,2005). TrkC-expressing neurons are mostly large cells (Wright and Snider,1995) which are in a direct contact with the skeletal muscle target (Stephens et al.,2005). In this study, the percentage of TrkA- and TrkB-expressing DRG neurons observed in both DRG cultures and neuromuscular cocultures is similar to that in DRG in vivo (Snider,1994; Wright and Snider,1995). The percentage of TrkC-expressing DRG neurons in DRG neuronal cultures observed in this study is lower than that in DRG in vivo (Snider,1994; Stephens et al.,2005). In neuromuscular cocultures of DRG neurons and target skeletal muscle cells, the percentage of TrkC-expressing DRG neurons is similar to that in DRG in vivo (Wright and Snider,1995) suggested that TrkC expression is partially dependent on the presence of target skeletal muscle cells.

In conclusion, our data suggest that target skeletal muscle cells are important for maintenance neurofilament phenotype but not neuropeptide phenotype and maintenance TrkC but not TrkA and TrkB expression in cultured DRG neurons. The expression of neurofilament and TrkC is regulated by target skeletal muscle cells is consistent with the close relationship between neurofilament- or TrkC-expressing DRG neurons and target skeletal muscle cells in vivo. Additional studies are necessary to clarify the mechanisms of interactions between neurofilament- or TrkC-expressing DRG neurons and target skeletal muscle cells.

LITERATURE CITED

  1. Top of page
  2. Abstract
  3. MATERIALS AND METHODS
  4. RESULTS
  5. DISCUSSION
  6. LITERATURE CITED