TRPV1 at nerve endings regulates growth cone morphology and movement through cytoskeleton reorganization


F. Hucho, Freie Universität Berlin, Institut für Chemie und Biochemie, Thielallee 63, 14195 Berlin, Germany
Fax: +49 30 83853753
Tel: +49 30 83855545

C. Goswami, Department of Human Molecular Genetics, Max Planck Institute for Molecular Genetics, Ihnestrasse 63-73, 14195 Berlin, Germany
Fax: +49 30 84131383
Tel: +49 30 84131243


While the importance of Ca2+ channel activity in axonal path finding is established, the underlying mechanisms are not clear. Here, we show that transient receptor potential vanilloid receptor 1 (TRPV1), a member of the TRP superfamily of nonspecific ion channels, is physically and functionally present at dynamic neuronal extensions, including growth cones. These nonselective cation channels sense exogenous ligands, such as resenifera toxin, and endogenous ligands, such as N-arachidonoyl-dopamine (NADA), and affect the integrity of microtubule cytoskeleton. Using TRPV1-transiently transfected F11 cells and embryonic dorsal root ganglia explants, we show that activation of TRPV1 results in growth cone retraction, and collapse and formation of varicosities along neurites. These changes were due to TRPV1-activation-mediated disassembly of microtubules and are partly Ca2+-independent. Prolonged activation with very low doses (1 nm) of NADA results in shortening of neurites in the majority of isolectin B4-positive dorsal root ganglia neurones. We postulate that TRPV1 activation plays an inhibitory role in sensory neuronal extension and motility by regulating the disassembly of microtubules. This might have a role in the chronification of pain.


cannabinoid receptor 1


dorsal root ganglia


growth cone-associated protein 43


isolectin B4








transient receptor potential, canonical type


transient receptor potential vanilloid receptor 1

Nerve endings and their growth cones are neuronal structures where microtubule dynamics and complex signalling events determine various biological functions. Though the importance of Ca2+ in such functions is established to a certain extent, the role of individual Ca2+ channels and the regulatory function of Ca2+ are not clear [1–6]. Recently, members of the transient receptor potential, canonical type (TRPC) channel family were reported to be present at growth cones and to play an important role in axonal guidance [7–12]. However, the mechanisms by which members of the TRPC family regulates axonal guidance is not clear.

Transient receptor potential vanilloid receptor 1 (TRPV1), a nonselective cation channel, is involved in pain signalling. TRPV1 detects various noxious physical and chemical stimuli resulting in influx of Ca2+. A high level of TRPV1 expression was reported in dorsal root ganglia (DRG) [13]. However, the distribution of TRPV1 with respect to tissue is under debate. Among the neuronal tissues, TRPV1 was also detected in many parts of the brain and spinal cord [14–16], suggesting that TRPV1's distribution is widespread and not restricted to peripheral neuronal structures.

Previously, we reported that the C-terminus of TRPV1 interacts with tubulin, and provides stability to microtubules under certain conditions [17]. However, we also demonstrated that activation of TRPV1 results in rapid disassembly of dynamic microtubules [18]. Based on these observations, we hypothesized that TRPV1 may regulate some specific neuronal functions where fine regulation of microtubule dynamics is important. In this work we extend our observations and demonstrate that TRPV1 is physically and functionally present at dynamic neuronal extensions, and their growth cones. Using DRG-derived F11 cells, embryonic DRG explants and dissociated DRG neurones, we show that TRPV1 modulates the cytoskeletal organization of neurite extensions and regulates neurite morphology and extension.


TRPV1 is localized in the central and peripheral zones of growth cones

Growth cones, the specialized structures at nerve endings, are known to regulate axonal growth. We explored if TRPV1 is localized at growth cones. As F11 cells reflect many properties of DRG neurones [19], we used this cell line as a model to express TRPV1. This cell line offers a better neuronal environment suitable for monitoring the localization and function of TRPV1 (therefore, in the following, we term the neurite-like extensions of F11 cells as ‘neurites’). Previously, we also used this cell line to explore how TRPV1 and the neuronal cytoskeleton are interrelated [17,18,20,21]. After transient expression in F11 cells, TRPV1 was detected intracellularly, at the plasma membrane, and throughout the neurites (supplementary Fig. S1).

By immunofluorescence analysis, we could detect TRPV1 at neurite endings that extend into growth cone-like structures. By co-immunostaining for the growth cone-associated protein 43 (GAP43/neuromodulin), a well-characterized growth cone marker [22–25], we confirmed the identity of these structures as conventional axonal growth cones (Fig. 1A).

Figure 1.

 TRPV1 regulates growth cone movements. (A) Localization of TRPV1 at a growth cone developed from a TRPV1 transiently transfected F11 cell. A merged image of TRPV1 (green) and GAP43 (red) is superimposed on the phase contrast image. The central zone (C), the transition zone (T) and the peripheral zone (P) of the growth cone is indicated. Black arrows indicate filopodial structures. Majority of anti-TRPV1 immunoreactivity appears at the C- and P-zone. (B) Examples of growth cones developed from TRPV1-GFP transiently transfected F11 cell. Live cell images reveal the presence of TRPV1-GFP (green) predominantly at the P- and C-zones of growth cone. Scale bar = 10 µm. (C) Time-lapse confocal images of a growth cone (the same one as shown in (B); see also supplementary Video S1) just after adding RTX. The red line and the black arrow indicate the borderline and the centre of the main shaft of the growth cone, respectively, at time 0. The blue arrow indicates the position of the main shaft at the centre of the growth cone at a particular time point during the retraction phase. In the last frame, the green arrow indicates a further extension of the main shaft.

Extending growth cones display distinct structural features, a central zone (C-zone), a peripheral zone (P-zone) and a transition zone (T-zone), which correlate with the presence of different cytoskeletal proteins [26–28]. The C-zone of growth cone is known to be enriched with vesicles of various sizes and with microtubules, in particular their plus ends. In contrast, the P-zone contains the plasma membrane and the actin cytoskeleton beneath it. In extending growth cones we observed a differential distribution of TRPV1 in these areas (Fig. 1A). Negligible TRPV1 immunoreactivity was detected in the T-zone whereas a significant TRPV1 immunoreactivity was observed in the P- and C-zones (Fig. 1A). To confirm further that this differential localization of TRPV1 at growth cones is not appearing during fixation, we expressed TRPV1-GFP in F11 cells and performed live cell imaging. We observed a similar distribution of TRPV1-GFP at the growth cones, i.e. TRPV1 is mainly present in the P- and C-zones (Fig. 1B).

Activation of TRPV1 results in retraction of growth cones

The importance of Ca2+ channels in the regulation of growth cone movements has been reported previously [1–6]. Earlier, we observed that activation of TRPV1 has a destabilizing effect on microtubules in the cell body [18]. Therefore, we investigated if activation of TRPV1 can also alter or regulate the cytoskeleton in growth cones and neurite extensions. For that purpose we expressed TRPV1-GFP in F11 cells.

We observed that the majority of the TRPV1-GFP positive growth cones responded to the application of the agonists resiniferatoxin (RTX) or capsaicin, causing retraction and/or collapse of growth cones. We observed that these cones (56 out of 60), which are dynamic, immature, not connected to each other and have an extending morphology, retracted quickly (Fig. 1C; see also supplementary Video S1). This process of retraction, due to TRPV1 activation, could be completely blocked by the antagonist 5′-iodoresiniferatoxin (5′-IRTX, data not shown), indicating that the retraction of growth cones was indeed due to TRPV1 activation. This not only indicates that TRPV1 can regulate the growth cone motility, but also suggests that TRPV1 activation leads to a rapid change in the cytoskeleton organization. This result was further confirmed with a true neuronal system (see later).

Activation of TRPV1 results in disassembly of microtubules and thus induces the formation of varicosities

We observed that in contrast to the above-mentioned growth cone, the extending cone at the end of long axon-like neurites does not retract upon TRPV1 activation, but rapidly turns to a pause state (growth cones at pause exhibit a distinct morphology) [28]. Their neurites develop multiple varicosities (not shown). Formation of varicosities is believed to represent a change in the microtubule cytoskeleton within neurites [29,30]. To confirm that TRPV1 activation mediated varicosity formation in long neurites is indeed due to disassembly of microtubules, we transiently expressed TRPV1 in F11 cells and compared the status of microtubules within neurites from TRPV1 activated and nonactivated cells. In the absence of activation, immunoreactivity against β-tubulin subtype III appears continuous along the neurites of TRPV1-expressing cells. In contrast, upon activation, it was no longer visible as a continuous pattern and often appeared as distinct discontinuous accumulations within the varicosities. The majority of these varicosities reveal TRPV1 immunoreactivity, indicating that these neurites indeed developed from transfected cells (supplementary Fig. S2a). In contrast, nontransfected F11 cells reveal a normal and continuous immunoreactivity for tubulin and do not form varicosities even after addition of RTX (supplementary. S2b).

Functional TRPV1 is expressed in embryonic (murine) DRG explants and are located at growth cones and neurites

To confirm that activation of TRPV1 results in similar changes not only in transfected F11 cells but also in a truly neuronal system, we used DRG explants from E12 and E13 mouse embryos. At this stage (E13) of development, we detected TRPV1-specific mRNA only in DRGs, but not in the spinal cord (Fig. 2A). However, we could not detect any specific TRPV1 immunoreactivity in these DRG explants by immunostaining (see Discussion). We presumed that expression of TRPV1 at this stage of development might be too low to be detected by immunostaining. Therefore, we probed the mice DRG explants by means of more sensitive methods, such as Ca2+-imaging, in response to pharmacologically specific components. We observed that some, but not all Fluo-4-AM loaded neurites and growth cones showed an increase in Ca2+ intensity after adding RTX (Fig. 2B). The increase in the Ca2+ influx was simultaneous throughout the ‘responding neurites’ and correlates very well with the formation of multiple varicosities all along the neurites (supplementary Video S2).

Figure 2.

 Functional TRPV1 is expressed in murine DRG neurones at embryonic stage E12. (A) RT-PCR analysis of the mRNA isolated from meurine E12 dorsal root ganglia (DRG, middle lane) and spinal cord (SC, last lane). (B) Functional TRPV1 is present in growth cones and neurites. Mouse DRG explants (E12) loaded with Fluo 4-AM was treated with 50 nm RTX. Fluorescence images before (a) and 2 min after RTX application (b) are shown in false colour (scale shown at right). ‘Responding’ neurites are indicated by arrows and nonresponding neurites by arrowheads. Significant TRPV1 activity is observed at growth cones and regions forming varicosities. Scale bar = 20 µm. (C) Mouse DRG explants (E12) loaded with fluo 4-AM were treated with 1 µm NADA. Fluorescence images before (a) and 20 min after (b) NADA application are shown. An enlarged (indicated by a white box) image of a NADA-responding neurite is shown in right (c). Scale bar = 100 µm.

As both RTX and capsaicin are exogenous natural agonists, we tested if endogenous ligands of TRPV1 can cause similar changes. We tested N-arachidonoyl-dopamine (NADA), which is present in neuronal tissues and is as potent as capsaicin [31]. We observed that application of NADA also resulted in influx of Ca2+ in some but not in all growth cones and neurites (Fig. 2C). Response to NADA (indicated by the arrow) is observed at the growth cone, throughout the neurites and also at neurite branching points. This confirms the presence of functional TRPV1 in ‘responding’ neurites. This observation is in agreement with the report that TRPV1 is expressed in the embryonic stage E13 ([32], also see Discussion).

TRPV1 activation leads to the collapse of murine growth cones and formation of varicosities

As embryonic DRG explants expressed functional TRPV1, we tested if activation of TRPV1 can also affect the microtubule cytoskeleton. Under control conditions all growth cones revealed a normal extended phenotype, and no neurites with varicosities were found (a total of 437 neurites from five explants were counted) (Fig. 3Aa,b). As expected, we observed that application of RTX to the murine DRG explant revealed distinct changes of growth cones and neurites (Fig. 3Ac,d). In response to RTX, 36.54% of growth cones (129 out of 353) revealed a ‘collapsed’ morphology, and neurites developed many varicosities (Fig. 3Ac,d; see also supplementary Video S3). Tubulin immunoreactivity was discontinuous in the responding neurites and was found only in the varicosities. However, nonresponding growth cones remained unaffected and retained normal neurites, even in the presence of RTX.

Figure 3.

 Activation of TRPV1 by RTX results in growth cone collapse and varicosity formation. (A) Activation of TRPV1 by RTX causes formation of ‘varicosities’ due to disassembly of the microtubule cytoskeleton. Neurites and growth cones originating from E12 mouse DRGs are shown (left side). Enlarged areas (indicated by white boxes) of the corresponding explants are shown at right side. Untreated explants (top panel, a and b), treated with agonist RTX in absence (middle panel, c and d) or presence of the antagonist IRTX (lower panel, e and f) are fixed and stained for tubulin. Arrowheads indicate neurites with collapsed growth cones and discontinuous immunoreactivity for tubulin at the varicosities. Scale bar = 20 µm. (B) Chicken DRG explants are insensitive to RTX, and do not form varicosities. The control (a) and RTX-treated (b) E7 chicken DRG explants are shown. Scale bar = 20 µm.

To prove that this phenotype is specific for TRPV1 activation, we preincubated the DRG explants with 5′-IRTX, the antagonist of TRPV1. Application of RTX in these 5′-IRTX preincubated explants resulted in the formation of varicosities or growth cone collapse with only 0.9% (only four out of 433, from a total of five DRG explants from mice) of the neurites (Fig. 3Ae,f). This not only proves that some neurites developed from embryonic DRG explants at this stage (E12) are sensitive to RTX, but also proves the functional presence of TRPV1 there.

Activation of a naturally occurring mutant form of TRPV1 does not result in the formation of varicosities

To confirm further the specificity of the TRPV1 activation-mediated varicosity formation in a different system, we used the DRG explants from E7 chicken as a negative control. It is known that the chicken homologue of TRPV1 does not respond to capsaicin or RTX due to a point mutation at the region of the capsaicin-binding site [33]. We observed that DRG explants isolated from chicken developed many growth cones within a day (Fig. 3B). Upon treatment with the agonist RTX, DRG explants from E7 chicken revealed a normal phenotype, and formation of varicosities was not observed at all (Fig. 3B). This demonstrates that the observed growth cone collapse and varicosity formation in mice DRG explant is indeed due to TRPV1 activation by RTX.

TRPV1 activation-mediated varicosity formation is partially Ca2+ independent

Next we tested the influence of direct Ca2+ influx on the TRPV1 activation-mediated varicosity formation. To test this, we added EGTA to the culture media before activation of TRPV1 with RTX. EGTA treatment alone does not result in any varicosity formation (Fig. 4Aa,b). However, we observed that in a medium with low Ca2+ (2 mm EGTA), application of RTX resulted in the formation of varicosities (Fig. 4Ac–d). However, in this condition, the tubulin immunoreactivity between two varicosities appears as a thin connecting line. This indicates that disassembly of microtubules was not complete and suggests that a partial conservation of microtubules occurs. Even at 20 mm EGTA, activation of TRPV1 by RTX caused varicosity formation (Fig. 4Bc,d). However, tubulin immunoreactivity between two varicosities again appeared as a thin connecting line. This indicates that while TRPV1 activation by RTX leads to varicosity formation, this process is partially independent of Ca2+ influx (see Discussion).

Figure 4.

 RTX-mediated varicosity formation is partly Ca2+ independent and due to the activation of TRPV1 located at the cell surface. (A) Varicosity formation is partly independent of Ca2+ influx. Mouse DRG explant treated with 2 mm EGTA for 15 min (a) and an enlarged section of the same (below, b) is shown. A DRG explant pre-treated with 2 mm EGTA for 15 min followed by activation with 50 nm RTX (c) and an enlarged section of the same explant (below, d) is shown. All explants are immunostained for tubulin and the arrowheads indicate the thin connection between two subsequent varicosities. Scale bar = 20 µm. (B) Tubulin immunostaining of a mouse DRG explant treated with 20 mm EGTA (a) is shown. A similar explant further treated with 50 nm RTX is shown on the right (b). Scale bar = 20 µm. (C) RTX-mediated varicosity formation is independent of Ca2+ influx from internal Ca2+ stores. Mouse DRG explants are treated with either thapsigargin alone (1 µm, 15 min) (left side, a) or with thapsigargin for 15 min and subsequently with RTX for 15 min (right side, c), then fixed and immunostained. Tubulin immunostaining of the explants (top panel, a and c) and an enlarged section (indicated by white box) of the corresponding explants (bottom panel, b and d) are shown. Thapsigargin treatment does not prevent TRPV1 activation-mediated varicosity (indicated by arrow) formation. Scale bar = 20 µm.

We observed that depletion of Ca2+ from the endoplasmic reticulum by incubating the explants with thapsigargin (1 µm for 35 min), a specific inhibitor of endoplasmic reticulum Ca2+-ATPase, does not cause varicosity formation (Fig. 4Ca,b). In contrast, addition of 50 nm RTX for 15 min to cultures that are preincubated with 1 µm thapsigargine led to the formation of varicosities (Fig. 4Cc,d). This indicates that varicosity formation is independent of Ca2+ mobilization from the endoplasmic reticulum.

NADA, an endogenous ligand of TRPV1, regulates growth cone morphology and motility

As application of NADA results in specific influx of Ca2+ in some neurites, we tested if NADA can also regulate growth cone morphology and movement. We observed that application of NADA also resulted in collapse of growth cones in murine DRG explants (Fig. 5A). Application of NADA also results in retraction of growth cones in the majority but not all neurites (supplementary Video S4). Some neurites were observed to also develop varicosities. However, NADA had a much weaker effect than RTX. Furthermore, NADA-mediated growth cone retraction is delayed, beginning about 20 min after application (supplementary Video S4).

Figure 5.

 NADA regulates growth cone morphology and movement. (A) NADA treatment (30 min) results in the collapse of a growth cone. Control E12 mouse DRG explants (a) and NADA-treated explant (c) were stained for tubulin (green) and actin (red). The majority of growth cones treated with NADA display collapsed morphology (b,d). Enlarged areas (white box) of the untreated (b) and NADA-treated growth cones (d) are shown inside. Scale bar = 100 µm. (B) Prolonged activation of TRPV1 by low level of NADA negatively affects the neurite extension of IB4-positive small-sized DRG neurones. Dissociated DRG neurones from adult rat were stained for neurone-specific β-III tubulin (green), IB4 (red) and DAPI. (a) Prolonged exposure with NADA (1 nm for 24 h) inhibits the growth of IB4 positive small-sized DRG neurones (indicated by arrows). Under this condition, IB4 negative neurones grow normally. Scale bar = 50 µm. (b) An example of an IB4-positive small DRG neurone grown under the above conditions. Note that these small IB4-positive neurones are fully viable in NADA-containing medium, even after 24 h of exposure, but develop very small neurites. Scale bar = 20 µm. (c) The majority of IB4 negative neurones do not respond to NADA (24 h). These neurones develop multiple extended neuritis, even in a medium containing 1 µm NADA. Scale bar = 50 µm. (d) In the absence of NADA, both IB4-positive and IB4-negative neurones grow normally and develop extended neurites. Scale bar = 50 µm.

As TRPV1 activation has a negative effect on neurite extension, we explored if prolonged but low-level activation of TRPV1 by NADA affects neurite extension and growth. Therefore, we tested the effect of a very low dose of NADA (1 nm) on dissociated DRG neurones. A significant fraction of TRPV1-positive small DRG neurones (>50%, by using tissue section) are isolectin B4 (IB4)-positive [34]. Therefore, in order to characterize the effect of NADA, we used IB4 as a marker for a subpopulation of DRG neurones.

We observed that this subpopulation of mainly small IB4-positive DRG neurones developed very small neurites when NADA was added for 24 h (Fig. 5Ba,b). No effect of NADA was observed in the majority of IB4-negative neurones under these conditions (1 nm) or even at higher concentrations (1 µm) of NADA (Fig. 5B). Under the same conditions, but in the absence of NADA, these IB4-positive, small neurones grow normally, develop many neurites and are comparable to the IB4-negative neurones (Fig. 5Bd). This indicates that in response to the endogenous ligand, TRPV1 can regulate both morphology and length of neurites in a subpopulation of DRG neurones.

In summary, TRPV1 seems to be present and functional in sensory neurones including growth cones at embryonic stages. Overall, our results suggest that TRPV1 regulates the microtubule cytoskeleton and affects growth cone morphology and motility.


Growth cones are well-organized dynamic structures involved in neurite pathway finding. Frequencies of Ca2+ waves generated by Ca2+ channels present in neurites and growth cones have a strong regulatory influence on neurite initiation, extension, branching and direction [1–6]. Here we provide evidence that TRPV1 is localized at specific areas of the growth cone and influences morphology and motility of neurites. We also show that the presence of TRPV1 at certain neurites and growth cones modulates the cytoskeletal reorganization by destabilizing them upon activation.

As the TRPV1 activation-mediated retraction of growth cones is fast (initiated within a few minutes) and results in disassembly of microtubules (18 and this study), it is unlikely that activation of TRPV1 will alter the ratio of the different cytoskeleton proteins. This suggests that the retraction of neurites is most likely to be independent of transcription or translation of a new gene product(s), but depends on the reorganization of structural proteins affecting both morphology and movement of neurites. Previously we had observed that activation of TRPV1 significantly affects the polymeric organization of the microtubule cytoskeleton. Activation of TRPV1 results in disassembly of dynamic microtubules but not of the actin cytoskeleton [18]. Growth cone retraction is a process involving both the actin and microtubule cytoskeletons. More specifically, the force generated by motor proteins is involved in this process [29,35]. We also observed that Taxol®, a microtubule cytoskeleton stabilizer, significantly delayed RTX activation-mediated varicosity formation, but could not block it completely (data not shown). Based on all these observations and previous evidence, we postulate that growth cone retraction and varicosity formation upon TRPV1 activation is a multi-step process that leads to sequential disassembly of microtubules (Fig. 6). In support of this hypothesis, we provide several lines of evidence. Firstly, after TRPV1 activation, immunoreactivity for tyrosinated tubulin (a marker for dynamic microtubules [36]) is diffusely present at the P- and T-zones of activated growth cones, including in the filopodial structures (supplementary Fig. S2C). This indicates a loss of dynamic microtubules due to TRPV1 activation. Secondly, long neurites frequently showed substantial formation of varicosities, a classic morphological symptom of microtubule depolymerization [29]. Finally, using embryonic DRG explants, we prove that activation of TRPV1 by RTX can result in formation of many varicosities along the neurites, most probably due to a very rapid disassembly of microtubules. However, some neurites are unchanged upon RTX application and do not develop varicosities. This suggests that TRPV1 is present in some but not in all neurites. This result is in agreement with the fact that a major fraction of DRG neurones is TRPV1 negative. Interestingly TRPV1 is not exclusively present at the growth cones of the ‘responding neurites’. For example, RTX application results in rapid increase in Ca2+ influx throughout the ‘responding neurites’ (Fig. 2), indicating that TRPV1 is also present all along the neurite shaft.

Figure 6.

 Proposed model of growth cone motility regulation by TRPV1. (A) Both anterograde force from the microtubule cytoskeleton (blue, upside arrow) and a retrograde force provided by actin cytoskeleton (red, down side arrow) determine the net axonal growth and movement. (B) TRPV1 activation-mediated growth cone retraction and varicosity formation is dependent on the status of the microtubule cytoskeleton. Stage 1: Activation of TRPV1 (indicated by arrow) results in partial disassembly of microtubules, leading to the retraction of growth cone. Stage 2: Further disassembly of microtubules leads to further retraction and initiates varicosity formation. Stage 3: Complete disassembly of microtubules results in a stage where further retraction is no longer possible. Stage 4: Retrograde force from actin cytoskeleton and complete disassembly of microtubules results in the varicosity formation.

Using time-lapse microscopy, we observed that upon RTX application, responding neurites first retract for a short time and then develop varicosities (supplementary Video S3). In most cases, within a single neurite, varicosity formation was almost instant and did not occur one by one, suggesting a simultaneous global change all over the neurites and a dynamic instability. This is in agreement with our previous observation that TRPV1 activation by RTX results in rapid disassembly of dynamic microtubules, but not of the actin cytoskeleton [18]. During the morphological transition from smooth neurites to varicosities due to RTX application, we frequently observed a short retrograde movement of the growing varicosities (supplementray Videos S1 and S3). This short retrograde movement within the retracting neurite suggests the involvement of functional actin cytoskeleton.

Growth cones are capable of recognizing different external stimuli and of integrating different cues in terms of Ca2+ transients [37]. Our data also indicate that chemical stimuli specific for TRPV1 can regulate the morphology and movement of TRPV1-positive growth cones. We demonstrate that NADA, an endogenous ligand of TRPV1, not only affects growth cone morphology and movement, but also the neurite length (Fig. 5B). Apart from NADA, there are few other putative endogenous ligands, such as N-oleoyldopamine known to activate TRPV1 [38]. Some other endovanilloids and fatty acid metabolites are also known to act on TRPV1 [39]. We did not test all of these endogenous ligands because they are not selective for TRPV1 and are also subjected to metabolic changes. NADA is also known to activate cannabinoid receptor 1 (CB1), although at higher doses [31,40,41]. Therefore involvement of CB1 receptors in the NADA-mediated process can not be ruled out completely.

TRPV1 activation by other noxious physical stimuli like low pH and high temperature are also expected to exert a similar phenotype. The combination of different physiological stimuli might produce a synergistic effect on the growth cone retraction and varicosity formation through TRPV1, but this is difficult to prove due to the presence of many other receptors sensing low pH and temperature changes. In agreement with our results, the presence of TRPV1 was recently detected at varicosities within embryonic spinal cords [42]. In this regard, it is important to mention that varicosities are often considered as synaptic terminals. Therefore, it might be possible that TRPV1 is involved in the synapse formation too.

Growth cone retraction and collapse due to RTX application is fast and seems to be irreversible, as addition of 5′-IRTX shortly after RTX application does not result in any recovery of growth cones. In contrast to RTX-mediated growth cone collapse and retraction, the NADA-mediated effect in primary neurones is much weaker and delayed. However, NADA exerts more physiological effects without affecting the cell's viability. The growth cones and neurites remain motile in NADA-containing medium, even 1 h after application. Using time-lapse imaging, we observed that after NADA treatment, retracted growth cones remained motile and started to re-grow after a short period of time (data not shown). Moreover, we could also show that at low concentrations of NADA (both with 1 nm and 1 µm), the majority of the IB4-positive dissociated DRG neurones survive for 24 h or more, but in these cases, the neurites grow very little (Fig. 5B).

We observed that growth cone retraction in response to TRPV1 activation is dependent on the nature and amount of the ligand. Strong agonists like RTX cause retraction for a short duration and subsequent formation of many varicosities, indicating that disassembly of microtubules is rapid, complete and most likely to be irreversible. In contrast, an endogenous ligand such as NADA causes retraction for a longer time but rarely forms varicosities. Application of NADA most probably results in incomplete and slow disassembly of microtubules. Thus effects of NADA are most likely to be reversible and physiological.

Our work also establishes another important aspect: TRPV1 is of functional importance in a subpopulation of DRG neurones already at an early developmental stage (E12–13 in mice). Recently, expression of TRPV1 has also been reported at embryonic stage (E13) by Funakoshi et al. [32]. Using tissue sections from embryonic mice, Funakoshi et al. [32] demonstrated that TRPV1 is present in certain nerve fibres. In contrast, we could not detect specific TRPV1 staining in our explant cultures or in dissociated DRG neurones by immunofluorescence analysis. These differences in the immunostaining might be due to the different antibodies used. However, it seems that the expression of TRPV1 in embryonic DRG neurones is subjected to different external conditions and is reduced upon culturing. Nevertheless, our results indicate that expression of TRPV1 at early embryonic stages can regulate the growth cone motility in certain neurones. This might have a developmental role, especially in response to endogenous ligands like endovanilloids, compounds belonging to the lipoxygenase pathway [31,38,39]. Further research must be conducted to confirm this aspect of TRPV1.

It is known that Ca2+ has a depolymerizing effect on microtubules in vitro[43,44]. This effect of Ca2+ on microtubules has also been demonstrated in vivo, and it has been shown that Ca2+ leads to two distinct processes: the ‘dynamic destabilization’ (a direct depolymerizing effect of Ca2+ on microtubules) and ‘signal-cascade-induced fragmentation’ of microtubules [45]. In the present work, we have attempted to determine the influence of Ca2+ ion and/or Ca2+-signalling on the TRPV1 activation-mediated varicosity formation. We demonstrate that varicosity formation and growth cone retraction is dependent on TRPV1 activation, but partly independent of Ca2+ influx. For example, RTX application results in varicosity formation even in the presence of 20 mm EGTA. We have demonstrated that varicosity formation is independent of Ca2+ release from the endoplasmic reticulum and involves TRPV1 receptors located at the plasma membrane. Additionally, we have observed that in TRPV1 transfected F11 cells, microtubule disassembly due to RTX application at low temperatures does not result in disassembly of microtubuli (data not shown). This may suggest the requirement of an additional step(s) (other than a direct effect of Ca2+) involving enzymatic or metabolic activity. Although the microtubule cytoskeleton acts as a downstream effector of TRPV1, the exact molecular mechanism underlying the changes due to TRPV1 activation remains to be determined.

Growth cones have a remarkable ability for chemoattraction and chemorepulsion. Recently, several members of TRPC channels were reported to be present at growth cones, and to regulate growth-cone morphology and/or functions [7–12,46]. For example, TRPC channels are important for brain-derived neurotrophic factor and netrin-1-induced growth cone turning [9,10]. Xenopus TRPC1 (XTRPC1) is also involved in growth cone motility and turning in response to growth factors [11]. The TRPC5 receptor has been reported to control neurite length: its activation leads to a decrease in the length of neurites [12,46]. Though we did not test the effect of different growth factors and all the known endogenous ligands on the TRPV1-positive growth cone movement, our results point to the similarities between TRPV1 and other TRPC channels, and indicate a common function of TRP channels in regulation of growth cone motility in response to endogenous compounds during axonal path finding.

In summary, our results suggest a functional implication of TRPV1 in the regulation of growth cone morphology and neurite movement. These may have relevance in some pathological and neurological disorders including chronification of pain.

Experimental procedures

Reagents and antibodies

RTX, capsaicin, 5′-IRTX, thapsigargin and Taxol® were purchased from Sigma Aldrich (Deisenhofen, Germany). NADA was purchased from Biomol (Hamburg, Germany). Mouse monoclonal anti-β-tubulin class III-specific antibodies (clone SDL.3D10), mouse monoclonal anti-α-tubulin specific antibodies (clone DM1A) and tetramethylrhodamine isothiocyanate-labelled IB4 from Griffonia simplicifolia plant were purchased from Sigma Aldrich. Rat monoclonal antibodies YL1/2, often used as microtubule marker was purchased from AbCam Ltd (Cambridge, UK). Rabbit polyclonal anti-N-terminal TRPV1 Ig were purchased from Affinity Bioreagents (Golden, CO, USA). Mouse monoclonal antibodies against neuromodulin, commonly known as growth cone-associated protein 43 or GAP-43 (clone 31) were purchased from BD-transduction Ltd (Heidelberg, Germany). Calcium sensor dye fluo 4-AM, Alexa-594-labelled phalloidin, Alexa-594-labelled anti-rat secondary IgG, Alexa-594-labelled anti-mouse secondary IgG were purchased from Molecular Probes (Invitrogen, Karlsruhe, Germany). Cy2-labelled-antigoat and Cy2-labelled anti-rabbit IgG were purchased from Dianova (Hamburg, Germany).


For heterologous expression in mammalian cells, the full-length rat TRPV1 cDNA subcloned in a pcDNA3.1 vector was used [17,18,20]. For expression of the C-terminally GFP-fused TRPV1, a cDNA fragment encoding rat TRPV1 was amplified by PCR using 5′-ATGGAACAACGGGCTAGCTT-3′ and 5′-TCTCCCCTGGGACCATGGAA-3′ primers and subcloned into pCDNA3.1/CT-GFP-TOPO vector (Invitrogen) [21].


RT-PCR amplification of DNA coding for a TRPV1-fragment was carried out using cDNA libraries separately isolated from DRG and spinal cord of embryonic mouse (E13). Specific primer sets (forward primer TGTACTTCAGCCA TCGCAAG and reverse primer CCAGGATGGTGATGG CTC) were used for amplifying a 534 basepair fragment of TRPV1. For control purposes, amplification of a natriuretic peptide receptor 3 fragment was carried out using the same cDNA isolated from DRG as well as from spinal cord.

Cell culture and transfection

F11 cells were cultured in Ham's F12 medium (Invitrogen) supplemented with 20% fetal bovine serum (Invitrogen). For transient transfection, lipofectamine (Invitrogen) was used.

TRPV1 activation assay and live cell imaging

For visualizing the effect of TRPV1 activation, F11 cells were seeded on glass cover slips. TRPV1 was expressed by transient transfection. Two days after transfection, F11 cells were incubated with Hank's balanced salt solution buffer at room temperature (25 °C) supplemented with 1 mm CaCl2 and RTX (100 nm) for 1 min, followed by immediate fixing with paraformaldehyde (PFA) (2%). Immunocytochemistry of fixed cells was performed as described previously [17,18,20,21]. For live cell imaging, F11 cells transiently expressing TRPV1-GFP were maintained in complete medium and RTX (100 nm) and Ca2+ (1 mm) was added. Antagonist 5’-IRTX (1 µm) was used for blocking TRPV1. All live-cell images and fixed-cell images were taken with a confocal laser-scanning microscope (Zeiss Axiovert 100 M) with a 63× objective and analysed using the Zeiss LSM image examiner software.


For immunohistochemistry analysis, the cells were fixed with 2% PFA for 5 min and fixed cells were permeabilized with 0.1% triton X-100 in NaCl/Pi (5 min). The cells were subsequently quenched with 2% glycine in NaCl/Pi and the cells were blocked either with 5% normal goat serum or 5% bovine serum albumin in Tris-buffered saline Tween-20. The primary antibodies were used at the following dilutions: rabbit polyclonal anti-TRPV1 Ig (1 : 1000), rat monoclonal anti-tyrosinated tubulin Ig (1 : 1000), mouse monoclonal anti-α-tubulin Ig (1 : 1000) and mouse monoclonal anti-GAP-43 Ig (1 : 1000). All primary and secondary antibodies were incubated for 1 h at room temperature in NaCl/Pi supplemented with 0.1% Tween 20 buffer. Fluoromount G was used for mounting the cover slips.

Embryonic DRG explants

The effect of endogenous TRPV1 channel activation on growth cones was assayed following methods developed by Raper and Kapfhammer [47]. Briefly, explants dissected from mouse E12 or chick E7 DRGs were grown at 37 °C overnight on poly l-lysine laminin-1-1 (Tebu-bio, Offenbach, Germany) coated Petriperm dishes in DMEM (Invitrogen) supplemented with hNGF-β (20 ng·mL−1, Boehringer, Ingelheim, Germany). The next day, cultures were incubated with either 100 or 50 nm RTX for 20 min to activate TRPV1. As a control, cultures were preincubated with the antagonist 5’-IRTX (1 µm) for 10 min, followed by addition of RTX. In some experiments, explants were preincubated for 15 min with 1 µm thapsigargin, 2 mm EGTA, 20 mm EGTA, or 2 µm Taxol® before being treated with RTX. The cultures were fixed with 0.25% glutaraldehyde in NaCl/Pi and subsequently stained for α-tubulin. Cultures were analysed with an Axiovert135 fluorescence microscope (Zeiss).

Time-lapse imaging and calcium imaging

Time-lapse images (phase contrast) were captured with an Axiovert200M microscope (Zeiss) equipped with different filter sets, a heated stage maintained at 37 °C and with a constant CO2 source (5%). For calcium imaging, DRG explants were loaded with Fluo-4 AM (final concentration 5 µm) for 30 min. To avoid an excess of fluo-4 dye, the medium containing dye was replaced by fresh medium. The fluo-4-loaded explants then were grown in the fresh medium for at least 2 h to recover from the shock due to medium change, before they were used for time-lapse imaging. Images were captured using specific filter sets (excitation BP 450–490 nm, emission 515–565 nm) every 30 s with an UV pulse of 10 ms. Images were analysed for changes in intensity of Ca2+-mediated fluorescence using lsm-examiner software and converted into Rainbow colour.

Dissociated DRG culture

Dissociated rat DRG neurones were prepared from adult male rats (200–300 g) following a protocol adapted from that of Hucho et al.[48]. Rats were anesthetized with CO2. L1-L6 DRGs were removed, desheathed, pooled and incubated with collagenase (final concentration 0.125%, for 1 h at 37 °C), followed by a trypsin digest (final concentration 0.25%; for 7 min at 37 °C). Cells were separated by trituration with a fire-polished Pasteur pipette. Dead cells and debris were removed by centrifugation (Hettich Rotanta/T, 5 min at 100 g). Cells were resuspended in 12 mL of NeurobasalA/B27 media supplemented with l-glutamate (25 µm) and l-glutamine (0.5 µm). Cells were plated (0.5 mL·well−1) on polyornithine laminin-1-precoated glass coverslips (12-mm diameter) and incubated in 24-well plates at 37 °C in 5% CO2. The neurones were grown for a total of 48 h, and NADA was added to the culture 24 h after plating. Neurones were fixed with 2% PFA and stained for βIII-tubulin (1 : 1000) and IB4-lectin (1 : 5000 dilution). Images of the neurones were taken with a confocal microscope (Zeiss). Overlapping images covering the entire length of all neurites was put together manually by using adobe photoshop software (PC-WARE Information Technologies, Leipzig, Germany).

All animals were handled according to the EU directive 86/609/EWG for the protection of experimental animals, Germany.


We thank D. Krück for technical assistance, R. Jahnel for preparing the TRPV1 constructs, F. G. Rathjen for valuable input. Financial support by BMBF, Deutsche Forschungsgemeinschaft, Sfb515, and the Fonds der Chemischen Industrie is gratefully acknowledged. We thank T. B. Hucho and L. Goswami for their support and suggestions. CG is currently supported by the Max Planck Institute for Molecular Genetics, Berlin, Germany.