The detection of painful stimuli takes place at the nociceptive peripheral sensory nerve terminals in the skin or the viscera. These miniature structures (0.5 to in diameter, Fig. 1) are pivotal for the detection of noxious stimuli.1,2 Consequently, the comprehensive understanding of nociception and hence mechanisms of pathological pain can be achieved only by detailed exploration of nociceptive terminal function. However, not much is known about their properties because they are barely accessible by conventional electrophysiological methods. The saphenous skin–nerve preparation and ex vivo somatosensory system models developed by Reeh3 and Koerber and Woodbury,4 respectively, as well as other models using extracellular recordings from nerve fibers5–7 monitor axonal activity following activation of the terminals, in different conditions, but not the activity at the terminals themselves. Electrophysiological recordings and ion imaging from terminal axons provided essential information about currents, which underlie action potential generation8–10 and terminal signaling.11,12 Nevertheless, spatial constraints of the electrophysiological method make it unsuitable for studying biophysical properties of signal propagation along the cylindrical terminal and distal axon. Conventional imaging assays overcome these spatial limitations; however, they are lacking in sufficient temporal resolution crucial for exploring ion dynamics underlying action potential generation and propagation. Many molecular and biophysical aspects of nociception have been described using nociceptor cell bodies, situated in dorsal root ganglions (DRG) or trigeminal ganglions.13–16 The functional environment17–19 and geometry of terminals differ from that of cell bodies, and likely, their passive membrane properties, density, and specific repertoire of transducer and voltage-gated channels. Hence, detailed characterization of ion signal onset, kinetics, and magnitude, along the terminals and terminal axons, have yet to be achieved, albeit being highly valuable for understanding nociceptive physiology. Consequently, the basic questions in nociceptive physiology, such as where action potentials are generated and how they propagate along nonmyelinated tiny axons in normal and pathological conditions, still remain obscure.
Here, we introduce a new approach for high-temporal resolution optical recordings of ion dynamics from a single nociceptive neurite terminal in vitro. This approach allows us to monitor capsaicin-induced calcium () and sodium () dynamics at the terminal neurites and characterize their propagation along the distal neurites. Using this approach, we have demonstrated that the capsaicin-induced and signals change as they propagate from the terminal and along the neurite. We showed that capsaicin-induced calcium signals initiated at the terminals were not dependent on voltage-gated channels, whereas signals measured away and onward, toward the cell body, were faster, stronger, and at least partially mediated by voltage-gated channels.
Materials and Methods
All animal procedures were approved by the Ethics Committee of the Hebrew University. Five- to six-week-old male C57BL/6 mice were used.
Dorsal Root Ganglion Compartmental Cultures
The compartmented chamber dishes were prepared similarly to that described by Campenot et al.20 Briefly, poly-lysine- and laminin-coated dishes were scraped with a pin-rake ( width of pins; Tyler Research) to create a series of parallel tracks on the laminin coating, limiting neurite growth to the areas left with laminin, in parallel lanes. A droplet of 0.4% methylcellulose (Sigma) in DMEM (Biological Industries) was spread to cover the scratched region where the Teflon compartmentalized chamber is placed. Teflon compartmentalized chambers (Teflon dividers, Camp10 or Camp320, Tyler Research) were attached to the culture dish with silicon grease (Dow Corning), applied with a syringe grease applicator (CAMP-GLSS, Tyler Research). The central compartment was used for plating the cell soma, flanked by peripheral compartments, containing lanes in which neurites can extend [Fig. 1(a)].
DRG culture preparation: Adult mice were deeply anesthetized (3% isoflurane), and DRG were isolated and dissociated in a manner similar to that described previously,15 with the following modifications: dissected ganglia were placed in ice-cold DMEM (Biological Industries), pelleted, and reconstituted for enzymatic digestion in solution containing collagenase and dispase II (Roche) for 45 min. Cells were triturated in the presence of 50 U DNase I (Sigma) and centrifuged through 10% bovine serum albumin (Sigma). The cell pellet was resuspended in 1 ml of Neurobasal (Gibco) medium containing B27 supplement (Invitrogen), penicillin and streptomycin (Biological Industries), 2.5S NGF (Promega), and GDNF (Sigma). Cells were plated on to poly-lysine- () and laminin- () coated 35-mm glass bottom tissue culture dishes [World Precision Instruments (WPI)] in the middle compartment of the compartmented chambers [Fig. 1(a), see also Ref. 20].
On the day of seeding the cells (day 0), both chambers contained the above-mentioned Neurobasal media with 2.5S NGF; thereafter, the concentrations of NGF were altered to promote the sprouting neurites to grow toward the distal chambers as follows (cell bodies and peripheral compartments, respectively): day and ; day and ; and days 5, 8, and day and .
Cultures were maintained at 37°C in a humidified incubator containing 5% . The cultured DRG cell somata underwent viral coinfection on either of 8- to 10-days postseeding. Imaging experiments of neurite terminals were carried out on days 11 to 14 from seeding. Teflon dividers were removed prior to the experiment.
To measure the changes in intraterminal and intraneuritic concentrations (), the adeno-associated virus serotypes 1 or 6, carrying an expression cassette for the genetically encoded indicator (GCaMP6s) or the mRuby florescent protein indicator (under a CMV promoter; ELSC Virus Core Facility), were used. Cells were infected by the two viruses simultaneously.
Image data were processed by Neuroplex software (RedShirt Imaging) and analyzed using pCLAMP 10.2 (Molecular Devices) and OriginPro 9.0.0 (OriginLab Corporation).
Number of repeats in this study (’s) refers to the number of terminals examined in each experiment. In the imaging assay, each analyzed terminal was from a culture made from a different animal. For the imaging assay, reflects different terminals in different dishes, however, not all from different animals.
All analyses were performed on the processes in the same field of view.
We used a short imaging protocol of up to 5 s to avoid photo damage, which occurred during longer exposure, probably due to the recurrent excitation with high-intensity ultraviolet and blue excitation wavelengths used here.
Fluorescent excitation was performed with a 75 W Xenon arc lamp (Lambda DG4; Shutter Instruments) and a GFP filter set (Ex 480, Em 535, dichroic Lp 510; Chroma). An inverted microscope (Eclipse Ti; Nikon) equipped with a oil 1.4NA objective, Epi-Fl attachment, and perfect focus system was used. For bright field imaging and fluorescent still image acquisition, an Exi Aqua monochromator camera (QImaging) and Nikon elements AR software (Nikon) were used. Changes in intraterminal and intraneuritic fluorescence were acquired using a back-illuminated cooled CCD camera (NeuroCCD-SMQ; RedShirt Imaging), controlled by Neuroplex software (RedShirtImaging). In these conditions, pixel size was . This spatial resolution was sufficient for analyzing the signal propagation along terminals and distal neurites, even though in some locations the diameter of the structure (0.5 to ) could have been less than the camera resolution.
Images were acquired at (fps).
To measure changes in intraterminal and intraneuritic concentrations () in vitro, the cell-permeable indicator SBFI-AM (Molecular Probes) was used. Five-millimolar stock was prepared in dimethyl sulfoxide with 10% Pluronic F-127. Cells were incubated for 1 h with a final concentration of SBFI-AM in the culture media, followed by a 30-min rinse period. Images were acquired at 125 fps. Florescent excitation was performed using a CoolLed fluorescence excitation system, using a 380-nm LED and a Fura2 filter set (Ex: 380 nm, Em: 510 nm, dichroic Lp: 400 nm; Chroma). To compensate for time-dependent decrease in fluorescence, due to fluorophore bleaching, fitted linear curves derived from the first 500 ms of the acquired data were subtracted from the recorded data.
All imaging experiments were performed in standard external solution (SES) composed of 145 mM NaCl, 5 mM KCl, 2 mM , 1 mM , 10 mM glucose, and 10 mM HEPES ().
Capsaicin (300 nM) or vehicle (SES with ethanol) was focally puff applied (500 ms puff, 2 psi, PV820 Pneumatic PicoPump; WPI) by a pipette with 4 to resistance, when filled with SES. The application regime was commanded by a Digidata 1440 A/D interface (Molecular Devices).
In some experiments, the channel blocker, tetrodotoxin (TTX, ; Alomone Labs) together with a Na(v) 1.8 channel blocker, A803467 (; Tocris), were applied to bath for 10 min before the recordings.
We calibrated the dispersion profile of puffed substances by measuring changes in the sulfarhodamine () fluorescent intensity profile with distance from the pipette tip [Fig. 1(b)] according to
To measure changes in within a terminal at the highest possible resolution our conditions permit, the data were collected from sized regions of interest (ROIs) [1 pixel; Fig. 1(c)]. To examine changes in concentration propagating along the neurite and to improve the signal-to-noise ratio, the data were collected using Kernel smoother from sized ROIs (; Fig. 2), which were distributed every along the neurite’s terminal and processes, dividing the terminals into segments. All the data were analyzed from the ROIs delimiting the examined segments, and the size of the ROI was neglected when the actual distance was calculated. Therefore, the distance is represented as an estimated location rather than an absolute distance.
The time of onset of the signal was measured as the time at the onset of a clear sharp deviation from fluorescent base level measured locally, while analyzing the trace from its positive peak toward time “0.” All the values were subtracted from the value at the terminal tip () of the corresponding terminal.
Difference in time of onset between two adjacent segments, i.e., the delay in onset at one segment compared with the time of onset at the previous segment, were calculated according to
The fluorescence propagation velocity within segments was calculated according to
Maximum rate of rise of transients was calculated as at different locations along the neurite terminal. The values were normalized to the value of the terminal tip of the corresponding terminal and plotted as a function of distance from the terminal tip.
All averaged data are presented as the . Assessment of statistical significance of differences between means was performed with repeated-measures of analysis of variance (ANOVA), with posthoc Bonferroni, as found appropriate.
Results and Discussion
To study processing of nociceptive information directly at its most relevant location, with sufficient time and space resolution, we introduced an approach for ultrafast optical recording of cultured nociceptive processes. To that end, we used compartmented chambers [Fig. 1(a), see Sec. 2] that have been previously used to study neurites of DRG neurons.21–24 In these chambers, DRG cell bodies are prompt to grow neurites, and terminal processes can be easily identified and accessed [Figs. 1(a) and 1(c)].
We selectively stimulated these neurite terminals with a calibrated focal puff application [Fig. 1(b)] of capsaicin, an agonist of the noxious heat-sensitive transient receptor potential cation channel subfamily V member 1 (TRPV1) channel, expressed by nociceptive neurons.25 We placed the pipette, containing 300 nM capsaicin, about from the terminal tip [Figs. 1(a) and 1(b)] because at this distance, the application of vehicle did not induce changes in GCaMP6s fluorescence [Fig. 1(c); terminals] or detectable movement of the processes. First, we calibrated the dispersion profile of the puffed substances [see Sec. 2; Fig. 1(b)]. Our data show that the predicted concentration at the terminal tip is of the initial value [Fig. 1(b)], i.e., in our conditions, the concentration of capsaicin at the terminal is close to the half maximal effective concentration () of TRPV1 channels to capsaicin (26). This is an estimate of the peak concentration during a 500-ms puff, which dissipates in the medium when the puff ends. Additionally, considering that the minimal concentration of capsaicin to activate TRPV1 channels is ,26 the prominent decrease in concentration along the -axis [Fig. 1(b)] renders the concentration mostly ineffective once it reaches other parts of the fiber or adjacent terminals. Indeed, in all the performed recordings, we did not observe responses in neighboring terminal branches of the neurite.
We measured capsaicin-induced increase in at the neurite’s terminal process using the genetically encoded indicator GCaMP6slow [GCaMP6s, Fig. 1(c)]. We chose the slow variant of the GCaMP6 indicator due to its better signal-to-noise ratio and sensitivity (compared with other GCaMP6 variants27), which enabled the detection of weak fluorescent emission signals from fine neurite processes. To study transducer channel- and action potential-mediated changes in terminal , we optically recorded changes in terminal GCaMP6s fluorescent intensity at a sampling rate of 1000 fps. As it has been shown that the half width of the action potential recorded from the distal neurite of nociceptive neurons is ,8 this sampling rate should be sufficient to detect reflections of single action potentials. We achieved this using a oil 1.4NA Plan Apo objective, in combination with a fast acquisition back-illuminated CCD camera (see Sec. 2). Puff application of capsaicin, but not vehicle [Fig. 1(c)], led to prominent increase in GCaMP6s fluorescence, detectable at a spatial resolution of along the terminal tip [Fig. 1(c)].
Next, utilizing the high functional spatial resolution of our approach, which allows precise tracing of signal propagation along the terminal, we measured the amplitude, time of onset, and rise rate of capsaicin-induced signals along the terminal and terminal neurite [Figs. 2(b), 2(d)–2(f), black]. It is noteworthy that due to short acquisition time (see Sec. 2), we could not explore the mechanisms underlying the decay phase of the capsaicin-induced response at the neurite’s processes and, therefore, we focused our efforts on the rising phase of the response.
We analyzed the difference in time of onset between two adjacent segments, consecutively for all adjacent segments (see Sec. 2), which reflects the time it took the signal to travel along a segment. We found that the signal traveled along for , for , and for (). After segment , time of onset difference between two ROIs was [segments to ; Fig. 2(d), black, ]. Using these data, we calculated the fluorescent propagation velocity of capsaicin-induced signals between segments (see Sec. 2) and demonstrated that the signal propagated along with a velocity of , with , and with (). Signals propagated between segments to did not vary in their response peak amplitude or maximum rise rate [; Figs. 2(b), 2(e), 2(f) black]. After (segment ), the fluorescence propagation velocity increased significantly and reached a peak of at (), and the speed of propagation between segments remained similar along the rest of the measured terminal (, one-way ANOVA, ). It is noteworthy that the values for propagation velocity that we have measured represent the propagation of fluorescent signals along the process and not the electrical conductance. The propagation of GCaMP6s fluorescence is dependent on electrical conductance; however, there are many additional factors that are involved, such as the channels’ and indicator kinetics (see also Ref. 28).
Remarkably, at (segment ), a two-phase response appeared [Fig. 2(b)]. While the first phase was similar to the one observed at the terminal tip ( for rise rate; for the amplitude, one-way ANOVA with posthoc Bonferroni), the second phase possessed a significantly larger peak amplitude and maximum rise rate [, one-way ANOVA, terminals; Figs. 2(e), 2(f) black], reaching a plateau of about 3.5 times larger response peak amplitude and 7 times larger rise rate at the maximum. This large and fast component was completely abolished following application of a combination of TTX and A803467 [Figs. 2(c), 2(d)–2(f) red] to block TTX-s- and Na(v)1.8-mediated TTX-r channels,29 which are expressed by nociceptive terminals.8 Moreover, blocking Na(v) channels delayed the time of onset [Fig. 2(d), red], reduced the fluorescence propagation velocity to , and decreased the peak amplitude and maximal rise rate [Figs. 2(e) and 2(f), red] of capsaicin-evoked responses recorded along the neurite after , such that the values become similar to those recorded at the first [Figs. 2(e) and 2(f)]. These data suggest that the increase in beyond the is partially dependent on activation of voltage-gated channels.
We further supported the latter notion by directly measuring capsaicin-induced fluxes in the neurite’s terminal process using the indicator SBFI-AM. Imaging of fluxes at different locations along the terminal revealed that application of capsaicin induced a small transient increase in , in the neurite’s terminal tip and a larger step-like signals at a distance from the tip [Fig. 3(b), ].
It is noteworthy to mention that TRPV1 channels are also permeable to .30 Thus, the signal at the terminal tip may reflect influx via active TRPV1 channels. The fast and step-like nature of the signals recorded away from the neurite may underlie burst-like firing of the action potentials.31 From a methodological point of view, this in vitro approach, which is based on a distinctive compartment containing only neurite terminals, is currently the only approach allowing direct study of terminal dynamics. It is advantageous over an in vivo assay as the usage of AM-based dyes in vivo will likely produce a strong background noise due to AM dye loading of not only the terminals but also nonneuronal cells in the imaged area. The lack of genetically encoded indicators for specific measurements of flux in distinct cell types render measurements from small structures in vivo or ex vitro extremely challenging. Importantly, low initial fluorescence intensity of SBFI-AM in the neurite’s terminal, due to multiple factors (e.g., relatively low passage of SBFI excitation wave length by the objective and intraterminal loading) allowed us to sample capsaicin-induced intraterminal changes of , only at 125 fps.
Altogether our data suggest that the nociceptive neurite’s terminals and processes, in terms of dynamics and its dependence on channel blockers, are composed of two functionally separated zones. Our results imply that signal generation and propagation in the first is not mediated by voltage-gated channel-induced depolarization. This signaling could be mediated by -permeable TRPV1 channels, low threshold voltage-gated channels, -induced release, and capsaicin-mediated release from TRPV1-expressing internal stores.32 Given the focality of our application and the susceptibility of the late response to channel blockers, TRPV1-mediated release from internal stores has a lesser contribution to the capsaicin-induced response, which we observed after . Hence, as suggested by our data, signal propagation beyond the first is largely driven by voltage-gated channels leading to elevation probably via voltage-gated channels and -induced release. Interestingly, the maximal propagation velocity, we have measured from terminal neurites of nociceptive processes, was substantially smaller than those measured using single fiber recordings from mice sciatic-tibial nerve in vivo.6 This discrepancy could result from the differences between electrical conductance and fluorescent propagation, as stated above. Indeed, the velocities of fluorescence propagation that we and others28 have measured in cultured neurites are slower than that measured for propagation of electrical signals.6
In this study, we have analyzed the activity of individual terminal neurites, which are easily identified and traced. It is difficult, however, to trace and record individual neurite activity along its full length because proximal neurites in culture create dense bundles of neurites.20,24 Utilization of our approach on individually labeled neurons, via micropipette injection of a spectrally different dye, may overcome this difficulty and provide essential information about the calcium dynamics along different compartments of peripheral processes.
In conclusion, we have introduced an approach for studying nociceptive terminal processes by bringing together and utilizing fast and sensitive optical recordings of ion dynamics from easily accessible cultured neurite terminals. Altogether, the data shown here demonstrate that our methodology gains in sensitivity and temporal acquisition capabilities, compared with standard imaging techniques, allowing fast optical recordings of weak signals from minute nociceptive terminal processes. Moreover, our approach gains in functional spatial resolution, compared with single electrode electrophysiological approaches, permitting characterization of signal propagations along the terminal and terminal neurites by sampling the signals in action potential relevant time resolution. Thus, the ultrafast and high-resolution optical recording technique described here provides the only tool for a detailed study of a nociceptive terminal functional molecular network, which underlies noxious stimuli detection and transmission in normal and pathological conditions. This could advance the very much needed knowledge for understanding pain physiology and pathophysiology. Moreover, this platform can be utilized to enable studying the terminals of other neuronal types, such as other primary sensory neuron terminals.
Support is gratefully acknowledged from the Deutsch-Israelische Projectkooperation program of the Deutsche Forschungsgemeinschaft (DIP) grant agreement BI 1665/1-1ZI1172/12-1 (RHG, BK, SL, and AMB); European Research Council under the European Union’s Seventh Framework Programme (FP7/2007-2013)/ERC grant agreement no. 260914 (RHG, BK, SL, and AMB); the Jacob and Lena Joels Chair for Excellence in the Life and Medical Sciences (AMB); and the Hoffman Leadership program (RHG). We would like to thank Dr. Maya Groysman, the manager of ELSC viral core facility, for designing and providing viruses used in this study.
Robert H. Goldstein received his BMedSc degree, magna cum laude, from the Hebrew University, Faculty of Medicine. He is currently a PhD candidate in neurobiology, under the supervision of Dr. Alexander Binshtok. The majority of his PhD work is focused on studying ion and voltage dynamics in peripheral sensory free nerve endings, in vivo and in vitro, both in normal and pathological states.
Ben Katz received his BSc degree in mathematics and philosophy. He moved to biological research and received his PhD in medical neurobiology entitled: “Structural and molecular dynamics in Drosophila photoreceptors,” under the supervision of Professor Baruch Minke. He continued to a postdoc training in medical neurobiology under the supervision of Dr. Alexander Binshtok, all of which were carried at the Hebrew University. His current research is studying a unique mutation in human TRPV1 channel.
Shaya Lev is currently working as a research associate and lab manager at Hebrew University. He received his PhD in physiology from the Hebrew University, Jerusalem, Israel, in 2010. He has published around 15 papers in different journals and conference proceedings. His research areas include neuro-cellular imaging, and cellular and molecular mechanisms of TRP channels, pain and nociception.
Alexander M. Binshtok is a PI at the Hebrew University Medical School studying cellular and molecular mechanisms of normal and pathological pain. He received his BSc degree in physical therapy from Ben Gurion University in 1995. His graduate work, with Dr. Michael Gutnick at the Hebrew University focused on cellular and molecular characteristics of cortical neurons. He did his postdoctoral fellowship with Dr. Clifford Woolf at Harvard Medical School, where he focused on developing approaches for pain-selective anesthesia.