The adrenomedullary chromaffin cell transduces chemical messages into outputs that regulate end organ function throughout the periphery. At least two important neurotransmitters are released by innervating preganglionic neurons to stimulate exocytosis in the chromaffin cell—acetylcholine (ACh) and pituitary adenylate cyclase activating polypeptide (PACAP). Although PACAP is widely acknowledged as an important secretagogue in this system, the pathway coupling PACAP stimulation to chromaffin cell secretion is poorly understood. The goal of this study is to address this knowledge gap. Here, it is shown that PACAP activates a Gαs-coupled pathway that must signal through phospholipase C ε (PLCε) to drive Ca2+ entry and exocytosis. PACAP stimulation causes a complex pattern of Ca2+ signals in chromaffin cells, leading to a sustained secretory response that is kinetically distinct from the form stimulated by ACh. Exocytosis caused by PACAP is associated with slower release of peptide cargo than exocytosis stimulated by ACh. Importantly, only the secretory response to PACAP, not ACh, is eliminated in cells lacking PLCε expression. The data show that ACh and PACAP, acting through distinct signaling pathways, enable nuanced and variable secretory outputs from chromaffin cells.
Introduction
The adrenal medulla is a core effector of the sympathetic stress response in the periphery (Cannon, 1940). When activated, it secretes epinephrine, norepinephrine, and important vasoactive peptides into circulation (Wolf et al., 2016). These hormones modulate cardiac, pulmonary, and metabolic functions in ways that favor survival or preserve internal conditions when they are likely to be disturbed (Cannon, 1940; Goldstein, 2010; Goldstein and Kopin, 2007).
Secretion from the adrenal medulla is dependent on input from preganglionic, sympathetic (splanchnic) fibers which terminate on adrenomedullary chromaffin cells (De Robertis and Ferreira, 1957; Grynszpan-Winograd, 1974). That the chromaffin cell secretory response is triggered by synaptically released acetylcholine (ACh) that has been appreciated for many decades. But whether ACh alone is sufficient to sustain catecholamine release, particularly under conditions where the adrenal medulla is strongly activated, is not as clear. Indeed, prior studies have revealed that catecholamine secretion exhibits a dramatic time-dependent decline when medullae are continuously perfused with ACh or when splanchnic input is electrically stimulated at high frequencies (Bevington and Radda, 1985; Wakade, 1988), such a decline is not evident with low-frequency, electrical stimulation of the splanchnic nerve (Wakade, 1988).
In the last decade, evidence has emerged that suggests pituitary adenylate cyclase activating polypeptide (PACAP), complements and, under specific conditions associated with strong sympathetic activation, supersedes ACh as the principal neurotransmitter at the splanchnic-chromaffin cell synapse (Eiden et al., 2018; Smith and Eiden, 2012; Stroth et al., 2013). PACAP was first identified in 1989 and described to cause an increase in cAMP in the hypothalamus (Miyata et al., 1989). Once PACAP became available as a pharmacological agent, it was shown to cause a long-lasting, non-desensitizing form of secretion from chromaffin cells (Przywara et al., 1996). Subsequent studies established the necessity of PACAP for sympathetic adjustments to insulin-induced hypoglycemia—a process regulated by adrenomedullary hormones—but evidence that PACAP caused chromaffin cell secretion, in situ, was lacking (Eiden et al., 2018; Hamelink et al., 2002; Smith and Eiden, 2012). This evidence was eventually provided by Smith and colleagues, who showed that high-frequency splanchnic stimulation of an adrenal slice fails to sustain catecholamine secretion when a PAC1R receptor antagonist, PACAP (6-38), is included in the slice bathing solution (Kuri et al., 2009; Smith and Eiden, 2012).
In comparison with ACh, the molecular mechanisms by which PACAP acts to stimulate secretion are poorly understood. The identity of at least part of the PACAP signaling pathway is encoded in its name; there is strong evidence that, in multiple cell types, exposure to PACAP leads to activation of adenylate cyclase and cAMP production leading to Ca2+ elevations (Blechman and Levkowitz, 2013; Dickson and Finlayson, 2009; Przywara et al., 1996). However, the PAC1 receptor is also known to couple to Gαq and phospholipase C β (PLCβ), which makes it unclear exactly where Gα and PLC couple receptor activation to exocytosis (Blechman and Levkowitz, 2013; Eiden et al., 2018). In addition, there is conspicuous lack of information on the properties of the Ca2+ signals that drive PACAP-stimulated exocytosis—information, which, if available, might provide deeper insights into the mechanisms by which intracellular Ca2+ is elevated. Finally, whether PACAP and ACh, operating through distinct intracellular effectors, differentially impact the fusion event itself or impinge on the fusion pore to regulate its expansion has not previously been investigated.
The goal of this study was to elucidate the mechanisms linking PACAP stimulation of chromaffin cells to dense core vesicle fusion. To this end, high-resolution imaging approaches were brought to bear on mouse adrenal chromaffin cells stimulated with PACAP or ACh and undergoing exocytosis. The data show that acute stimulation of chromaffin cells using PACAP elicits a form of exocytosis that requires signaling through a Gαs, Epac, and phospholipase C ε (PLCε). The Ca2+ transients that follow PACAP stimulation, imaged with a genetically encoded Ca2+ indicator, have a variable amplitude, do not show evidence of desensitization, and are sensitive to an inhibitor of L-type channels. Exocytotic events stimulated by PACAP and ACh also differed in their form and frequency. While the release profile of catecholamines from single vesicles was largely insensitive to the identity of the secretagogue, peptide release was slower in chromaffin cells stimulated by PACAP than by ACh. While ACh caused a burst of exocytotic events that waned in frequency as stimulation persists, the secretory response to PACAP did not run down, but persisted at a steady rate through the stimulation period. In chromaffin cells lacking PLCε expression, the secretory response to PACAP was eliminated. However, no deleterious effects on ACh-stimulated secretion in the PLCε KO were detected. These data suggest that PACAP and ACh regulate parallel and independent pathways for exocytosis in chromaffin cells. The nuanced and variable responses of cells to these neurotransmitters may be important for tuning adrenomedullary hormone release to the changing needs of an organism during stress.
Materials and methods
Animals
C57BL/6J mice (also referred to as wild-type mice herein) were obtained from Jackson Labs. PLCε KO mice were generated by Smrcka and colleagues (Wang et al., 2005). Animals were group housed with two to five per ventilated cage with a 12 h dark/12 h light cycle with full access to food and water. Animal procedures and experiments were conducted in accordance with the University of Michigan (PRO00007247 and PRO00009235), the University of Toledo (400138), and the Rowan University IACUC protocols (2017-039).
Mouse chromaffin cell preparation and transfection
Mice (2–4-mo old) were gas anesthetized using an isoflurane drop jar technique and sacrificed by cervical dislocation. For TIRF and electrophysiology experiments, three to four male or female mice were euthanized for two plates to ensure proper cell density and health of chromaffin cells. For Fura-2AM imaging experiments, male mice were used (see below). Adrenal glands were removed from mice and moved to dishes containing ice cold mouse buffer (148 mM NaCl, 2.57 mM KCl, 2.2 mM K2HPO4 3 H2O, 6.5 mM KH2PO4, 10 mM dextrose, 5 mM HEPES free acid, and 14.2 mM mannitol). Using a dissection microscope, the cortex was carefully removed using forceps (Cat.# 72891-Bx; Dumont Swissmade) and microscissors (World Precision Instruments, 14124-G). The medullae were rinsed three times in 75 μl drops of papain enzyme solution (450 units/ml Papain [#LS003126; Worthington Biochemical], 250 μg/ml BSA, and 75 μg/ml dithiothreitol) for 15 min at 37°C shaking 140 rpm in 0.5 ml of papain solution. After 15 min, the digesting solution was mostly removed and replaced by 0.5 ml of collagenase enzyme solution (250 μg/ml BSA, 0.375% collagenase, 0.15 mg/ml DNase). Digestion was continued for another 15 min at 37°C shaking 140 rpm. The digestion was stopped by transferring glands into an antibiotic-free culture medium (Dulbecco’s modified Eagle’s medium [DMEM]; Thermo Fisher Scientific) supplemented with 10% fetal bovine serum (FBS; Thermo Fisher Scientific). The glands were then triturated by push-pull movement using a 1-ml pipette tip (up to eight times). The suspension containing glands was then spun at 1,300 × g for 2.5 min. Supernatant was discarded and the pellet was resuspended in 300 μl of antibiotic-free medium and triturated again in a 200-μl pipette tip (up to eight times). The suspension was spun again at 1,300 × g for 2.5 min. After discarding the supernatant, the pellet was re-suspended in 110 μl of resuspension buffer (Invitrogen, Thermo Fisher Scientific) for transfection. The desired plasmid was added to the mixture (7 μg/106 cells). The suspended cells were transiently transfected by electroporation with a single pulse (1,050 mV, 40 ms) using the neon transfection system (Invitrogen, Thermo Fisher Scientific). Before digestion step, 35-mm-diameter dishes with 14-mm-diameter glass-bottom dishes (#P35G-1.5-14-C; MatTek Corporation) were pre-coated with Matrigel (Cat.# 356230; Corning) diluted in DMEM (1:7) for 1.5 h, after which the dishes were washed with DMEM. After electroporation, 200 μl of antibiotic-free medium was added to cells. 100 μl of electroporated cells were then deposited in each dish (two dishes total per condition). The cells were stored in an incubator (37°C, 5% CO2) for ∼4 h. Culture medium with antibiotics was then added to a final volume of 2 ml DMEM supplemented with 10% FBS, 9.52 unit/ml penicillin, 9.52 µg/ml streptomycin, and 238 μg/ml gentamicin (Thermo Fisher Scientific). The media was changed the day after plating and experiments were usually conducted 24–48 h after plating.
Neuropeptide Y (NPY) constructs (in pEGFP-N1) were provided by Dr. Ronald W. Holz (University of Michigan, Ann Arbor, MI). The Lck-GCaMP5G plasmid was obtained from Addgene (Cat.# 34924). The PLCε-FLAG-P2A-mCherry insert was synthesized and cloned into a pCMV-script EX vector (Genscript). For rescue experiments in the PLCε KO chromaffin cell in which PLCε-FLAG was overexpressed, transfected cells were identified based on their mCherry fluorescence.
Patch-clamp electrophysiology
Patch-clamp pipettes were made from 8250 glass (King Precision Glass). Data were recorded using an Axopatch 1C patch-clamp amplifier (formerly Axon Instruments; now Molecular Devices). Data acquisition was performed using custom data acquisition routines (donated by Stephen R. Ikeda, National Institutes of Health/National Institute on Alcohol Abuse and Alcoholism, Rockville, MD) within Igor Pro software (WaveMetrics) on a Macintosh minicomputer with an InstruTech ITC18-USB data acquisition board (HEKA Elektronik/Harvard Bioscience). Voltage recordings were made in the perforated patch, current-clamp configuration using 70–100 µg/ml of amphotericin B (Millipore-Sigma) backfilled into the recording pipette, sampled at 5-kHz, digitized, and stored on the computer for later analysis. All experiments were performed at room temperature. Data analysis was performed using Igor Pro software (WaveMetrics) and GraphPad Prism.
For current clamp recordings, the external (bath) solution contained (in mM) 140 NaCl, 5.3 KCl, 10 HEPES, 0.8 MgCl2, 2 CaCl2, and 15 glucose, pH 7.4. The pipette solution contained (in mM) 135 KCl, 8 NaCl, 2 MgCl2, and 20 HEPES, pH 7.2 (osmolality, 300 mOsm/kg). Pipettes were front-filled with this pipette solution and back filled with the same solution supplemented with 70–100 µg/ml of amphotericin B. Solutions containing ∼1 µM PACAP (Tocris Bioscience/R&D Systems) or 100 μM acetylcholine (Millipore-Sigma) were applied locally to cells using a Warner Instruments VC-6 perfusion system (Harvard Bioscience). The measured series resistance at the start of each recording was 40 ± 4 MΩ (n = 40). In every cell, a 5-mV step was given every 10 s in voltage clamp mode until reasonable access was attained, which generally took from 15 to 30 min. Cells were then switched to current clamp mode to make the recordings.
Ratiometric imaging of cytosolic [Ca2+] in FURA-2AM loaded cells
Methods are similar to those previously reported (Brindley et al., 2016). A piece of coverslip with mouse chromaffin cells attached was washed twice with HEPES-buffered Hanks’ balanced salt solution and incubated with 5 μM Fura-2AM (Setareh Biotech, LLC) at 37°C for ∼45 min, protected from light. Cells were then washed in Fura-free solution for 30–60 min at room temperature, protected from light. The coverslip was transferred to a recording chamber mounted on a Nikon TE2000 fluorescence microscope (Nikon Instruments Inc.) The recording bath was continually perfused with extracellular solution from gravity-fed reservoirs at a flow rate of ∼3 ml/min. Ratiometric imaging was performed using an InCyt IM2 fluorescence imaging system (Intracellular Imaging Inc.). Typically, between 4 and 8 individual cells were selected in the field of view for analysis. Cells were excited at wavelengths of 340 and 380 nm and emission detected at 510 nM with a PixelFly digital camera. The ratio of fluorescence emission at the two excitation wavelengths (340/380 nm) was measured every 1 s and converted to an estimated [Ca2+] using an in vitro calibration curve generated by adding 15.8 μM Fura-2 pentapotassium salt to solutions from a calibration kit containing 1 mM MgCl2 and known concentrations of Ca2+ (0–1,350 nM). In the presence of magnesium, the Kd of FURA is ∼220 nM (Grynkiewicz et al., 1985), although additional factors in the intact cell including pH, viscosity, protein concentration, and temperature can also impact Kd. Therefore, the reported concentrations are considered estimates rather than precise measures of [Ca2+]. Data are presented from 26 individual chromaffin cells recorded from 5 independent experiments and 2 separate cell preparations. Each cell preparation pooled cells from two male wild-type mice. The extracellular recording solution contained (in mM) 145 NaCl, 2 KCl, 1 MgCl2. 6 H2O, 10 glucose, 10 HEPES, and 2 CaCl2, pH 7.3, and osmolarity ∼305 mOsm. After a stable baseline, the bath solution was switched to one containing PACAP (100 nM) for 60 s. This resulted in a sustained rise in [Ca2+] that persisted even after washout of the PACAP. The cells were then sequentially exposed to 1 or 10 μM nifedipine to selectively block calcium entry through L-type voltage-gated calcium channels as shown in the representative trace (see Fig. 4). FURA2-AM was prepared as 1 mM stock solution in DMSO and aliquots stored for 1–2 wk at −20°C and protected from light. Nifedipine was prepared as a 10 mM stock in ethanol and stored and protected from light at −20°C until immediately before use.
TIRF microscopy
An Olympus cell TIRF-4line microscope (Olympus) was used to perform TIRF imaging. The microscope utilizes a TIRF oil-immersion objective (NA 1.49, 100×) with an additional 2× lens in the emission path between the microscope and electron-multiplying charge-coupled device camera (iXon Ultra 897, Andor Technology). The final pixel size of images was 80 nm.
All TIRF experiments were performed between 35 and 37°C. Physiological salt solution (PSS) buffer was prepared (145 mM NaCl, 5.6 mM KCl, 2.2 mM CaCl2, 0.5 mM MgCl2, 5.6 mM glucose, and 15 mM HEPES, pH 7.4) and prewarmed to 37°C before experiments were performed. Culture medium was replaced with warmed PSS before the experiment and changed after each stimulation protocol. Chromaffin cells were individually stimulated using a needle (100 μm inner diameter) connected to a perfusion system under positive pressure ALA-VM4 (ALA Scientific Instruments). For experiments in which ACh and PACAP were applied sequentially, cells were exposed to basal PSS for 5 s and then stimulated with 100 μM ACh for 60–90 s, washed with basal PSS for 30 s, and then stimulated with 1 μM PACAP for 90–120 s. Otherwise, cells were individually stimulated by local perfusion of either ACh or PACAP for an equivalent 60–90 s period. ESI-09 (Cat.# 4773; Tocris) was added to the bath at the concentration indicated in the Results for at least 10 min prior to stimulation with agonist. 8-(4-Chlorophenylthio)-2′-O-methyladenosine-3′,5′-cyclic monophosphate acetoxymethyl ester (cpTOME-AM; Cat.# 4853; Tocris) was locally perfused onto chromaffin cells at the concentration indicated in the Results. Where applicable, cells were incubated with 100 µM FFN511 (Cat.# ab120331; Abcam) in imaging buffer for 1 h at 37°C. FFN-loaded cells were washed with PSS prior to imaging on the TIRF microscope.
Experiments using pharmacological agents were always performed on the same day/prep as experiments on untreated, control cells. Experiments on PLCε KO chromaffin cells were always performed on the same day as WT cells or PLCε KO cells in which PLCε was overexpressed (rescues). Only data from same-day controls are shown in Figs. 3, 7, and 9. For Fig. 2, data from multiple experimental weeks were pooled and are presented together.
Image analysis of fusion events
Fusion events for cells containing either NPY-pHluorin or FFN were identified visually. Regions of interest (ROIs) were selected using the Time Series Analyzer v3.0 plugin on Fiji software. Manually selected fusion events were selected using ROIs with diameters measuring 800 nm. The intensity of fluorescence was measured for each frame for each ROI. A nearby ROI, where no fluorescent punctum was evident, was used for background subtraction.
For the experiments where the secretion event is characterized by a precipitous loss of fluorescence from an ROI that includes pre-secretion fluorescence-marked granules in its field (e.g., all FFN511 release events), we use a custom-written IDL program called “droprate” that characterizes (in a largely unbiased, hands-off manner) the rate of the loss (i.e., the dropoff). The goal of the analysis of fluorescence vs time data for “dropoff” responses is to: (1) locate the time of the dropoff event and (2) quantitate the rate of the decline around that time. These two functions are performed entirely automatically and identically for each run to avoid introducing bias. The main problem to be overcome by the analysis is random noise. As an aid to understanding, Fig. 1 follows the effect of each mathematical analysis step on an actual run.
The program automatically analyzes the original fluorescence vs. time data Forig(ti) (shown in Fig. 1) as follows:
Time of dropoff
Forig(ti) (Fig. 1, black) is smoothed by a running boxcar average, with a boxcar width of 30 time bins on either side of each time point ti. Within that width, the points are all equally weighted in the average. The purpose is to suppress high-frequency noise that otherwise might obscure or misidentify the midpoint of the drop-off time. This boxcar averaging is performed three successive times. The result is Fsmooth(ti) (Fig. 1 A, yellow). We are interested in where Fsmooth(ti) drops most rapidly, or equivalently, where the running difference Fdiff(ti) = Fsmooth(ti-1) − Fsmooth(ti+1) reaches a maximum (Fig. 1 A, light blue). The time location of that maximum is defined as the time tdrop at the steepest part of the dropoff (Fig. 1 A, red).
Rate of decline
Once tdrop is found, we no longer use Fsmooth(ti) for any calculations. Instead, we revert to the original Forig(ti) to evaluate the rate (i.e., the steepness) of the decline around that time. A time window centered around tdrop, with a constant width that is much broader than any dropoff event in any of the runs is defined (dark blue lines in Fig. 1 A and zoomed-in in Fig. 1 B). Forig(ti) within that window normalized and shifted in amplitude to vary from +1 at the beginning of the window to −1 at the end (This normalization and shift is only provisional; the data will soon be least-squares fit by a mathematical shape whose beginning and end values and the slope at tdrop will be adjustable as free parameters). That windowed, time-and-amplitude adjusted (but unsmoothed) original data are called Fwindow (Fig. 1 B, black).
By a standard least-squares fitting procedure, the theoretical Ffit(bt') (Fig. 1 B, green) is fit to the experimental Fwindow, with a, b, and c as free adjustable parameters. The parameter b (in s−1) is the reported rate of the dropoff.
Dropoff rates returned by this procedure are valid only up to about the reciprocal of the sample time of the experimental data (0.018 s here). For example, if some hypothetical original data are noise-free and drops cleanly from +1 to −1 in only one time bin, its (hypothetical) dropoff time (= b−1) could be as long as the sample time or as short as 0. In that extreme case of truly short dropoff times, the fitting method used above will return a dropoff time that does not exceed = b−1 but could be even less.
In contrast to the FFN511 dropoff events described above, NPY-moxGFP release events often displayed distinct rise and fall phases (see examples in Fig. 5). The time duration of NPY release was calculated using a different custom analysis program written in IDL called “secrate” (previously described in: Abbineni et al., 2019; Bendahmane et al., 2018; Bendahmane et al., 2020; Bohannon et al., 2017). Briefly, in this program, the user defines the start and end times in the fluorescence-versus-time curve for each event. The value of the fluorescence at the chosen start time tstart, just before fluorescence begins to rise, is considered the baseline. The end time tend is chosen to be where the fluorescence after the event has returned to its lowest value. The program then determines the time of the maximum fluorescence tmax within this time window. The intervals (tstart, tmax) and (tmax, tend) are defined as the rise phase and fall phase, respectively. Each of those phases is first best-fitted with a fifth-degree polynomial (for smoothing), and then a weighted average slope is calculated for each interval, upward for the rising phase and downward for the falling phase. Straight lines with those slopes are then pinned to the positions of the maximum (upward or downward) slopes of the fluorescence data and extrapolated to the baseline. The time period between the baseline intercept of the rising phase straight line and the falling phase straight line is considered to be the duration of the event.
In all cases, the release durations measured by the fitting algorithms were compared to visually determined release durations for accuracy. Events from multiple cells were pooled for analysis.
Image analysis of Ca2+ signals
Cells were transfected with Lck-GCaMP5G (Akerboom et al., 2012; Shigetomi et al., 2010) for qualitative analyses of Ca2+ changes upon ACh or PACAP stimulation. ROIs were manually selected and the equation %ΔF/F was used to calculate fluorescence changes within each ROI. In qualitative terms, ACh invariably produces a large, sudden, and early onset fluorescence increase in the defined field of view, then monotonically and smoothly decaying thereafter. PACAP produces a more variable and smaller response (sometimes none at all), with individual “sparks” exhibiting a longer onset delay at seemingly random times, and a non-monotonic course that is “noisy” at a wide range of time scales. This noise is, in part, likely due to different distinct sparks occurring at different times and locations with the field of view.
In quantitative terms, we separate each trace into two phases: the ACh phase and the subsequent PACAP phase, by computer-marking the time (t = 0) at the beginning of the ACh trace and at the border transition time between the ACh and PACAP phases. The absolute exact time of that border mark is occasionally different for different runs. We then quantified the following features for each of the two phases, for each run, with exactly the same protocol deployed by a custom-written IDL program called “calspike”: (a) the maximum amplitude; (b) the number of maxima; and (c) the time duration of each maxima. Because the curves are often quite noisy (due to shot noise and to the superposition of individual signals at different times), measurement of each of these parameters requires some pre-analysis data preparation, as follows.
Maximum amplitude. Because of noise, there are many more “maxima” than genuine fluorescence spikes. Therefore, we smooth the data by convoluting it with an exponential decay “kernel.” This smoothing is mathematically equivalent to a low-pass filter with a Lorentzian shape in the frequency domain. The 1/e width of the kernel is set at 50 time bins (approximately equal to 2.5 s here). The smoothing convolution is performed twice in succession. The kernel itself is adjusted by subtracting off its mean so that the kernel itself has a zero mean. That ensures that the very-low-frequency components of the smoothed trace are eliminated (i.e., the baseline is zeroed-out). What is left is a smoothed curve without baseline drifts that nonetheless preserves the positions and amplitudes of any spikes in the data. Then, the amplitudes of all the surviving maxima are recorded.
Number of maxima. From these same smoothed and baseline-flattened curves, the number of maxima can be counted.
Time duration of a spike. Often, the two minima surrounding each distinct maximum do not have the same amplitude. In other words, the signal from a typical spike (or set of cotemporaneous spikes) does not return to the same level before another spike appears. This makes the question of determining the width of a spike a bit more complicated. Starting with the smoothed, baseline flattened curves, we draw a straight, often sloping, line between the two minima surrounding a maximum. The value of that straight line at the time of its enclosed maximum is the new “baseline” of that maximum. Then, the position of a horizontal line at a designated constant fraction β (=0.9 here) of the amplitude from that baseline value to the maximum amplitude is determined. The full width of the maximum is defined as the time difference between the two crossings of that horizontal line with the smoothed data (one crossing on the upslope, the other on the downslope).
Western blotting
Adrenal glands were dissected from 2–4-mo-old mice and transferred to dishes containing ice-cold mouse buffer (148 mM NaCl, 2.57 mM KCl, 2.2 mM K2HPO4.3H2O, 6.5 mM KH2PO4, 10 mM dextrose, 5 mM HEPES free acid, and 14.2 mM mannitol). Adrenal medullae were obtained from those glands by trimming off the surrounding adrenal cortex. Both the medullas (left and right) were combined and lysed using hand-held homogenizer in an Eppendorf tube carrying 1× RIPA buffer with protease inhibitor. Lysate were further sonicated for two pulses, 15 s each at 20% amplitude. Samples were then centrifuged at 5,000 × g for 5 min at 4°C. Supernatant was transferred to a fresh tube and protein concentration was measured using BCA assay. Equal amounts of protein were loaded on a gradient (4–12%) NuPAGE Bis-Tris gel and separated in 1× NuPAGE MOPS SDS running buffer for 2 h at 100 V (room temperature). Proteins were then transferred to a PVDF membrane in 1× NuPAGE transfer buffer for overnight at 120 mA. Membrane was blocked with 5% milk in TBST for 1 h at room temperature. Membrane was washed 3× with TBST for 5 min each and primary antibodies were added and incubated at 4°C for overnight. Membrane was washed again with 3× with TBST for 5 min each. HRP-conjugated secondary antibodies were added and incubated at room temperature for 2 h. Blots were then developed using an ECL reagent in an iBright imaging system. Antibody information is as follows: rabbit anti-PLCε, 1:5,000, Smrcka lab (in-house, clone 2163), rabbit anti-Epac2, 1:200, Santa Cruz Biotechnology (Cat.# sc-28326), rabbit anti-PAC1R, 1:500, Abcam (Cat.# ab183103), rabbit anti-Synaptotagmin-7, 1:500, Synaptic Systems (Cat.# 105173), rabbit anti-PACAP, 1:5,000, Abcam (Cat.# ab181205), and rabbit anti-alpha tubulin, 1:5,000, Cell Signaling Technology (Cat.# 2144S).
Cyclic AMP assay
Cyclic AMP production was measured using the AlphaScreen cAMP kit from Perkin Elmer as directed by the manufacturer. Data were collected on Varioskan LUX Multimode Microplate Reader. The assay was repeated on three separate occasions, and data combined.
Statistical analysis
GraphPad Prism was used for all statistical analysis. Curve fitting was performed on Origin 2020b or IDL as described in the Materials and methods. The distribution of values for a particular data set was first tested for normality with a Shapiro–Wilk test. Differences in means of normally distributed data were subsequently compared using a t test with Welch’s correction. Kolmogorov–Smirnov, Mann–Whitney, or Wilcoxon tests were used to compare data sets whose individual values were not normally distributed (Dwivedi et al., 2017). For multiple comparisons, either a one-way ANOVA (normally distributed data), Kruskal–Wallis, or Friedman’s test was used to compare data sets. All mean values are reported in figure legends ± SEM to two or three significant digits. Within figures, the symbol ns means P > 0.05, * means P ≤ 0.05, ** means P ≤ 0.01, *** means P ≤ 0.001, and **** means P ≤ 0.0001. All p-values are reported to one significant digit except in cases were P > 0.99. In specific instances, the P values of 0.01 (Fig. 4 F) or 0.001 (Figs. 9 F and 10 B) were only achieved by rounding down. Here, the number of asterisks reflects the higher, unrounded P values.
Results
PACAP- and ACh-stimulated Ca2+ signals have distinct features
In the adrenal medulla, the binding of PACAP to PAC1Rs on postsynaptic chromaffin cells stimulates a cascade of events culminating in Ca2+-triggered exocytosis. The major goal of this study was to elucidate the pathway linking PAC1R activation to increases in Ca2+ and membrane fusion, which, in chromaffin cells, remains poorly understood. To achieve this goal, we first characterized the consequences of PACAP stimulation on the levels of intracellular Ca2+. To this end, we transfected mouse chromaffin cells with the genetically encoded Ca2+ indicator, GCaMP5G, fused to the N-terminal domain of Lck (Akerboom et al., 2012; Shigetomi et al., 2010). Lck causes membrane targeting of GCaMP5G, greatly improving the signal within the region of the cell best imaged in TIRF. Transfected cells, identified by their dim, green footprint, were sequentially stimulated with 100 μM ACh and then 1 μM PACAP (Fig. 2 A). This approach enabled direct comparisons to be made between the effects of ACh and PACAP on intracellular Ca2+ signals.
ACh elicits a large Ca2+ transient that rapidly returned to baseline (Fig. 2 C); PACAP elicits signals that were much smaller in amplitude and follow a complex pattern that is highlighted both in the image series (Fig. 2 B) and the %ΔF/F versus time graph (Fig. 2 C and boxed inset). In this example, intracellular Ca2+ levels climb slowly to a peak upon PACAP application and then fluctuate as stimulation proceeds. Meanwhile, the pattern of the ACh-stimulated Ca2+ transient was remarkably consistent from cell to cell, closely resembling the example provided in Fig. 2 C.
To facilitate analysis of the intensity versus time records of the Ca2+ response within an ROI, an analysis program called calspike was employed (see Image analysis of Ca2+ signals). The program automatically identifies peak fluorescence amplitudes (Fig. 2 D), the number of maxima (Fig. 2 E), and the duration of maxima (Fig. 2 F) within each record. The location of automatically identified maxima, or peaks, is evident in the inset of Fig. 2 C as vertical-dashed lines. The number of ACh-stimulated Ca2+ peaks within an ROI was usually 1; the number of PACAP-stimulated peaks varied from cell to cell. In the example shown, many peaks are evident, most of which were automatically identified by the analysis program.
ACh-stimulated Ca2+ transients were larger in amplitude than those caused by PACAP (Fig. 2 D), which were not only small, but increased and decreased with a variable time course. Consequently, PACAP stimulation was associated with a greater number of fluorescent maxima and a shorter maxima duration compared to ACh (Fig. 2, E and F). We also evaluated the impact of ACh and PACAP co-application on Ca2+ signals. As shown in Fig. 3 A, the monotonically decaying signal characteristic of ACh dominates the fluorescent ΔF/F when the neurotransmitters were co-applied. The signal associated with PACAP-stimulation was buried within, rather than super-imposed upon, the fluorescent ΔF/F record (Fig. 3 A). Neither the amplitude nor the duration of Ca2+ spikes were different between cells co-stimulated by ACh and PACAP and those cells stimulated by ACh alone (Fig. 3, B and C).
The elevations in intracellular Ca2+ triggered by PACAP can arise from the influx of Ca2+ through voltage-gated channels, or from Ca2+ that is released from internal stores (Dziema and Obrietan, 2002; Tanaka et al., 1996). In mouse chromaffin cells, it has previously been reported that PACAP-stimulated Ca2+ elevations arise from the influx of Ca2+ through T-type, low-voltage-activated channels (Hill et al., 2011). To investigate the pathway for the PACAP-stimulated Ca2+ elevations, ratiometric imaging of chromaffin cells loaded with Fura-2AM was performed. Mouse chromaffin cells express several subtypes of voltage-gated calcium channels, including CaV1 (L-type) channels which account for 40–50% of the whole cell current, CaV2 channels (N-, P/Q-, and R-type) which account for the majority of the non-L-type current, and perhaps a small contribution from CaV3 (T-type channels) under certain recording conditions (Carbone et al., 2019; Garcia et al., 2006; Novara et al., 2004). Exposure of cells to 100 nM PACAP caused a long-lived increase in intracellular Ca2+ that was significantly reduced in a concentration-dependent manner by nifedipine, a selective blocker of L-type calcium channels (Fig. 4, A and B). When external Ca2+ was removed at the end of the experiment, intracellular Ca2+ levels returned rapidly to baseline. These data suggest that acute exposure of mouse chromaffin cells to PACAP causes elevations in intracellular Ca2+ that rely primarily on Ca2+ influx through L-type channels.
To interrogate the effects of PACAP on the excitability of the chromaffin cell, patch-clamp electrophysiology experiments were performed. The results are shown in Fig. 4, C–E. Chromaffin cells frequently exhibit spontaneous electrical activity, a property that has been extensively described by other groups (for review, see Lingle et al., 2018). The application of PACAP caused only a small depolarization of the membrane potential of 2.6 ± 0.63 mV (from a resting value of −56 ± 3.1 mV, n = 9) in non-spiking cells, and 2.9 ± 0.97 mV (from a resting value of −50 ± 0.71 mV, n = 10) in spiking cells. In these 19 cells, the mean voltage change induced by PACAP is significantly different from 0 (P = 4 × 10−5; Wilcoxon signed-rank test). In separate experiments, performed only in spiking cells, PACAP caused a significant increase in the frequency of spikes, from a mean of 1.2 ± 0.21 per second prior to PACAP application to 1.5 ± 0.22 per second during PACAP application (P = 0.01, Wilcoxon matched-pairs signed rank test; Fig. 4, F–G).
In contrast, ACh caused a much larger depolarization of the membrane potential (13.8 ± 2.2 mV; n = 8; Fig. 4 E). Sometimes, this depolarization was superimposed on a burst of spikes (see fourth ACh trace from the top, Fig. 4 C). In four of eight cells, a transient hyperpolarization followed the initial depolarization (see first ACh trace, Fig. 4 C). Similar effects of ACh on cultured rat chromaffin cells were previously reported by Kidokoro et al. (1982).
Peptides hormones are released slowly from PACAP-stimulated fusion pores
The major consequence of cell stimulation by PACAP is secretion. Therefore, exocytosis was monitored in cells stimulated by PACAP and, for comparison, ACh. Mouse chromaffin cells were transfected with NPY-moxGFP to label secretory vesicles. The mox-variant of GFP is monomeric and optimized for trafficking through the reactive compartments of the secretory pathway (Costantini et al., 2015). A sudden, stimulation-dependent disappearance of a fluorescent punctum on the chromaffin cell footprint signaled the location of an exocytotic event. Examples of events triggered by ACh and PACAP are shown in Fig. 5 A. The time course of the moxGFP fluorescence was used to calculate release duration (see Image analysis of fusion events). The distribution of NPY-moxGFP release durations from many cells is shown in Fig. 5, B and C. Overall, there was a small, but significant difference in the release duration of NPY between cells stimulated by ACh and PACAP. The efficacy of PACAP and ACh as secretagogues was also compared (Fig. 5 C). Because the exact number of fusion events in a chromaffin cell varied, and more fusion events might be expected to occur in a larger cell, the number of fusion events was normalized to the surface area of the cell footprint (Fig. 5 D). This analysis showed that, for an equivalent period of stimulation, ACh caused roughly five times more fusion events than PACAP.
Fig. 5 shows that vesicles fusing in response to PACAP stimulation release NPY more slowly than vesicles fusing in response to ACh stimulation. Does PACAP stimulation also cause smaller cargos, like catecholamines, to be released slowly? To evaluate this possibility, chromaffin cells were incubated with FFN511 to label secretory vesicles. FFN511 is a false neurotransmitter that is transported into vesicles by the endogenous chromaffin vesicle monoamine transporter, VMAT (Gubernator et al., 2009). FFN511 loosely mimics the topology and physical properties of the monoamine neurotransmitters (Gubernator et al., 2009). Therefore, the release time course of FFN511 should be similar to the release time course of the endogenous catecholamines, epinephrine, and norepinephrine.
Fast (60–80 Hz), single-color imaging of FFN511 release was performed on a TIRF microscope. Examples of individual release events are shown in Fig. 6, A and B. Individual durations for many fusion events releasing FFN511 are plotted in Fig. 6, C and D. Release durations were calculated using an automated analysis program (described in Materials and methods). FFN511 was discharged more rapidly from fused vesicles than larger peptide cargos (compare release durations in Fig. 5 B to Fig. 6 C). However, the release duration of FFN511 was similar whether cells were stimulated by ACh, PACAP, or ACh and PACAP together (Fig. 6 C). Thus, the fusion pores of PACAP-stimulated events are slowed at a late stage of expansion, after the release of FFN and, therefore, catecholamines has already occurred.
PACAP causes Ca2+ entry through a pathway that is dependent on PLCε
The PAC1 receptor has previously been shown to couple to Gαs and Gαq to produce its cellular outputs (Blechman and Levkowitz, 2013). Accordingly, the signaling cascades activated by PACAP vary depending on the subtype of receptor (PAC1R versus VPACR) and system studied (Eiden et al., 2018; Holighaus et al., 2011; Macdonald et al., 2005; Spengler et al., 1993). It was recently shown that in the chromaffin cell, the relevant pathway for PACAP-stimulated exocytosis involves cAMP and Epac signaling through the PAC1R, downstream of Gαs (Kuri et al., 2009). Indeed, pharmacological activation of the exchange protein activated by cAMP (Epac) with the Epac-selective cAMP analog, cpTOME, elicits a form of catecholamine release that is similar to direct PACAP stimulation (Kuri et al., 2009). To substantiate a pathway that involves cellular effectors of Gαs, and especially Epac, cpTOME was directly perfused onto chromaffin cells expressing Lck-GCaMP5G. The Ca2+ responses to cpTOME application were qualitatively similar to the responses measured after PACAP stimulation (Fig. 7, A and B). Instead of the characteristic single spike of fluorescence that follows ACh stimulation, cpTOME-stimulated Ca2+ responses fluctuate with a variable time course.
If PACAP signals through Epac to drive Ca2+ influx, inhibition of Epac activity should reduce PACAP-stimulated Ca2+ transients. To test this idea, chromaffin cells expressing Lck-GCaMP5G were incubated with 1 μM ESI-09—a membrane permeable Epac inhibitor (Almahariq et al., 2013)—prior to stimulation with ACh and PACAP. Two exemplar intensity versus time records for this experiment are provided in Fig. 7 C. The presence of ESI-09 does not appreciably disrupt the ACh portion of the response (Fig. 7 D). However, ESI-09 reduces the %ΔF/F amplitude and the number of maxima (i.e., peaks) measured in cells stimulated by PACAP (Fig. 7, E and F).
Epac activates Rap small GTPases, which, in turn, bind to and stimulate PLCε (Smrcka et al., 2012). PLCε was recently shown to have important roles in Ca2+-induced Ca2+ release downstream of β-adrenergic receptors in cardiomyocytes and GLP-1 receptors in pancreatic β cells, in addition to regulating insulin vesicle exocytosis (Dzhura et al., 2010; Dzhura et al., 2011; Oestreich et al., 2009; Oestreich et al., 2007). Based on these observations, it was proposed that Epac, Rap, and PLCε form a “signaling module” in cell types in which these proteins are expressed (Dzhura et al., 2010; Oestreich et al., 2009; Zhang et al., 2013). We next investigated a novel role for PLCε—and by extension, this signaling module—in chromaffin cell exocytosis. To this end, we utilized a transgenic mouse model in which PLCε expression has been disrupted (hereafter referred to as PLCε KO or simply KO; Wang et al., 2005). However, to first ensure that any changes in the responsiveness of PLCε KO cells to PACAP were due principally to the loss of PLCε and not other proteins essential for transducing the effects of PACAP; medullae harvested from PLCε KO were probed for expression of PACAP, PAC1R, and Epac2 (Fig. 8). Syt7 expression was also assessed because of its important role as a Ca2+-sensor for exocytosis in chromaffin cells (MacDougall et al., 2018). Protein expression (i.e., intensity) was normalized to expression of α tubulin. Overall, the relative abundance of these proteins in PLCε KO adrenals was similar to, or slightly greater than, WT adrenals (Fig. 8, B–F). Cyclic AMP production in dissociated WT and PLCε KO cells was also assessed using an AlphaScreen fluorescent assay that is sensitive to free cAMP (Zhang and Xie, 2012). Even in the absence of PLCε, there was no apparent disruption in cAMP production over a wide range of forskolin concentrations (Fig. 8 G).
Buttressed by the knowledge that important signaling components upstream of PLCε were intact, we next evaluated Ca2+ responses to ACh and PACAP in WT and KO cells (Fig. 9). All chromaffin cells, irrespective of PLCε expression, produced an immediate and rapidly desensitizing response to ACh (Fig. 9 C). However, in the absence of PLCε, the PACAP-stimulated %ΔF/F was greatly reduced (Fig. 9, C and F). Sporadic, small fluctuations in Lck-GCaMP5G fluorescence were sometimes detected in the KO, but only rarely rose above 10%ΔF/F (2 out of 14 cells). Conversely, ACh-stimulated Ca2+ signals were not deleteriously affected by the absence of PLCε (Fig. 9, G and H).
To ensure that the absence of a Ca2+ response to PACAP in the PLCε KO cell was due specifically to the loss of PLCε, rescue experiments were performed. A plasmid encoding Lck-GCaMP5G was co-transfected with another encoding PLCε-FLAG-P2A-mCherry. Cells expressing PLCε-FLAG were identified by their mCherry fluorescence (expression of PLCε-FLAG was verified post hoc). KO cells in which PLCε expression was rescued (KO + PLCε) exhibited normal responses to both ACh and PACAP stimulation (Fig. 9, B, D, and F).
PLCε is required for PACAP-stimulated exocytosis
PACAP-stimulated increases in intracellular Ca2+ were greatly reduced in PLCε KO cells (Fig. 9). Moreover, the lack of responsiveness of the PLCε KO cells to PACAP was not due to a reduction in PAC1Rs, Epac2, or cAMP production (Fig. 8). To determine whether PLCε is also necessary for normal secretory responses to PACAP stimulation, chromaffin vesicles were loaded with FFN511 and stimulated through sequential application of ACh and PACAP (Fig. 10). The fusion efficacy of chromaffin cells was measured by dividing the total number of fusion events detected during the period of stimulation, by the surface area of the footprint. As shown in Fig. 10 A, ACh-stimulated secretion was not dependent on PLCε expression. Conversely, the secretory response to PACAP was strongly disrupted in the PLCε KO (Fig. 10 B). Indeed, only 1 event in 10 cells was observed in KO cells stimulated by PACAP. The secretory response to PACAP was restored in KO cells by transfection of a plasmid encoding PLCε.
ACh and PACAP cause kinetically distinct secretory responses
A characteristic feature of the secretory response to ACh is that it runs down over time. This is evident in Fig. 10, C and D, in which the occurrence of a fusion event is indicated by a vertical line, as well as cumulative frequency histograms that display the overall distribution of fusion event times (Fig. 10, E and F). Fig. 10 E shows that the cumulative histogram of fusion times in cells without PLCε (black dashed line) is similar to the cumulative histogram of fusion times in WT cells (solid red line). Thus, fusion triggered by ACh was largely independent of PLCε. Note that while the portion of the curve associated with ACh stimulation was best fit by a single exponential function (red dashed line), the PACAP response was best fit by a regression line (blue dashed line; Fig. 10 F). In contrast to ACh, PACAP caused a secretory response that is slow but proceeds at a constant rate.
Recapitulation of major results
Here, we report that binding of PACAP to PAC1 receptors activates a Gαs-coupled pathway for secretion in chromaffin cells that signals through Epac and PLCε. Stimulation of PLCε activity is necessary for Ca2+ influx through L-type channels and subsequent exocytosis (Fig. 11). In contrast to ACh, PACAP elicits Ca2+ signals that are spatially heterogeneous, exhibit a complex pattern, and are variable in amplitude. Fusion events caused by PACAP release peptide cargos more slowly than those triggered by ACh. The overall secretory response to PACAP is also kinetically distinct from the response to ACh. Vesicles fuse at a slower rate when exposed to PACAP than ACh. However, fusion does not appreciably run down during the period of PACAP stimulation.
Discussion
The adrenal chromaffin cell transduces messages released by innervating sympathetic neurons into outputs that shape the properties of end organ function throughout the periphery (de Diego et al., 2008). These messages come principally in two forms—acetylcholine and PACAP (Guerineau, 2019). Acetylcholine—its chemistry, synthesis, breakdown, and mechanisms of action in the adrenal medulla—had been studied since at least the middle part of the 20th century (Cannon, 1940; Coupland and Holmes, 1958; Douglas, 1968). By comparison, PACAP is a relative newcomer to the domain of adrenomedullary biology. Its role as an important neurotransmitter of the splanchnic-chromaffin cell synapse has only been established within the past 20 yr (Carbone et al., 2019; Eiden et al., 2018; Hamelink et al., 2002; Joseph et al., 2015; Kuri et al., 2009; Smith and Eiden, 2012). Accordingly, while the key features of cholinergic transmission in the adrenal medulla are known and widely accepted, important aspects of the pathway by which PACAP stimulates chromaffin cell secretion are either ambiguous or unexplored.
The notion that ACh must collaborate with another “non-cholinergic substance” to regulate catecholamine release from the medulla was first convincingly demonstrated many years ago (Wakade, 1988). This was based on experiments which showed a pronounced, nicotinic receptor-mediated desensitization of the secretory responses when splanchnic input to the medulla was stimulated at high frequencies (e.g., 10 Hz; Wakade, 1988). Interestingly, if the stimulation frequency was suddenly lowered to 3 Hz, the secretory response quickly rebounded. This low-frequency-stimulated, tonic discharge of catecholamine from the medulla was shown to be insensitive to atropine and hexamethonium—therefore, not mediated by ACh—and “facilitated” by phorbol ester (Wakade, 1988). Given its known effects on chromaffin cells, PACAP is now presumed to be that non-cholinergic substance. What remains unresolved is whether PACAP’s major role in the medulla is to complement ACh-stimulated secretion, or whether it may supersede ACh as the primary driver of medullary secretion under specific conditions. On the one hand, there is convincing data that at higher rates of splanchnic activity, after nicotinic receptors are desensitized, PACAP can “take over” to maintain catecholamine secretion and sustain the stress response (Kuri et al., 2009; Stroth et al., 2013). However, such ideas appear to be at odds with the relative weakness of PACAP, compared to ACh, as a secretagogue in cultured cells and in the intact medulla (Chowdhury et al., 1994; Wakade, 1988). Thus, our understanding of PACAP’s effects on chromaffin cells, in vitro and in situ, is far from complete. This knowledge gap motivated our experiments.
General features of Ca2+ signals and exocytosis elicited by ACh and PACAP
Acute stimulation of chromaffin cells with PACAP cause small and variable elevations in intracellular Ca2+ (Fig. 2). In this respect, the features of the PACAP-stimulated Ca2+ signals in chromaffin cells resemble signals in neurons of the suprachiasmatic nucleus in terms of their quite remarkable heterogeneity (Kopp et al., 1999). ACh-stimulated Ca2+ signals were more spatially homogenous and larger in amplitude than those stimulated by PACAP. This pattern of Ca2+ increase may reflect a wider distribution and/or higher expression of receptors for ACh which, when activated, more strongly depolarize the membrane potential and broadly open Ca2+ channels. A proportionally lower expression of PAC1 compared to ACh receptors may be offset by a tighter clustering to channels and local production of signaling metabolites (i.e., cAMP) to still enable an efficient secretory response.
Although the amplitude of the Ca2+ response to PACAP was lower than the response to ACh—at least at the concentrations of ACh and PACAP applied here—secretion was slower to desensitize (Fig. 8). One suspects that if the duration of PACAP stimulation was further lengthened relative to ACh stimulation, PACAP-stimulated fusion events would eventually outnumber ACh-stimulated events. However, within a brief 1–2-min stimulation window, ACh was a more effective secretagogue than PACAP. This finding is consistent with what has been previously reported in rat chromaffin cells stimulated by ACh and PACAP (Chowdhury et al., 1994). Such differences permit us to speculate upon divergent functional roles of ACh and PACAP as drivers of adrenomedullary hormone secretion, in situ. Ostensibly, the secretory response to ACh is designed to be rapid and immediate while PACAP enables slower but sustained release.
It has been suggested that PACAP elicits Ca2+ entry by opening a low voltage-activated (LVA), T-type channel (Hill et al., 2011; Kuri et al., 2009). However, a role for the T-type channel has been questioned because these currents are barely detectable in the mouse chromaffin cell in which these experiments were performed (Garcia et al., 2006). Carbone and colleagues showed that T-type currents do emerge in rat chromaffin cells, but only after a multi-day incubation with pharmacological activators of the Epac signaling pathway (Novara et al., 2004). Therefore, the immediate secretory response to PACAP stimulation is more likely to involve a Ca2+ channel that is more abundantly expressed in the medulla. In this study, we find that PACAP drives Ca2+ influx through a pathway that is sensitive to dose-dependent inhibition by nifedipine—an L-type channel blocker (Garcia et al., 2006). While roles for other channels were not evaluated, much of the PACAP-stimulated Ca2+ response was eliminated after nifedipine was applied to cells (Fig. 4 A). This explanation seems plausible, as CaV1 currents are known to be enhanced downstream of Gαs activation in both cardiac and neuronal cells (Dolphin, 2018; Qian et al., 2017). At the same time, because not all the Ca2+ signal was eliminated by nifedipine, it seems possible that PACAP ultimately stimulates secretion by activating multiple Ca2+ sources.
When characterizing the effects of ACh in this study, we did not attempt to differentiate between those mediated by nicotinic versus muscarinic receptors; its major conclusions did not depend on us doing so. Although it is often assumed that secretion stimulated by ACh occurs primarily through nicotinic receptors, muscarinic receptors are also coupled to a secretory response in a species-specific manner (for review, see Olivos and Artalejo, 2008). For example, muscarinic stimulation of bovine cells stimulates phospholipase synthesis and increases intracellular Ca2+ through a pathway involving IP3 (Fisher et al., 1981). However, no appreciable secretion is observed. Evidently, the amount of Ca2+ mobilized in this way is insufficient to elicit exocytosis (Cheek and Burgoyne, 1985). On the other hand, stimulation of muscarinic receptors in the rat and guinea pig adrenal causes mobilization of Ca2+ from caffeine-sensitive intracellular stores to a level that enables exocytosis to occur (Guo et al., 1996; Guo and Wakade, 1994; Ohta et al., 2001). Both nicotinic and muscarinic receptors also contribute to the ACh-stimulated secretory response in mouse chromaffin cells (Calvo-Gallardo et al., 2016), with muscarinic effects being mediated by a PLC (Matsuoka and Inoue, 2017; Olivos and Artalejo, 2008). Note that neither the Ca2+ signals nor secretion stimulated by ACh was disrupted in the PLCε KO. Thus, if there is a major muscarinic contribution to the ACh-stimulated secretory response, it must also operate independently of the epsilon isoform.
Effects of PACAP on cellular excitability
In patch-clamped cells, PACAP stimulation caused only a small, 2–3 mV depolarization of the resting membrane potential (RMP; Fig. 4 E). The RMP of these mouse chromaffin cells was usually at or near −50 mV, which is similar to values previously reported for chromaffin cells maintained in culture (for an extensive review on this topic, see de Diego et al., 2008). Perfusion of PACAP onto chromaffin cells did cause an increase in frequency of action potentials. These action potentials were “spontaneous” in many patch-clamped cells, occurring at a rate of ∼1.2 Hz. When PACAP was applied, action potential frequency increased significantly to 1.5 Hz (Fig. 4 F). Thus, we find that the major effect of PACAP perfusion was not to cause a large shift of the membrane potential to a more depolarized value as ACh frequently does (Fig. 4 E), but rather to increase the excitability of the cell in a more subtle way. These subtle effects on cellular excitability leave open the important question of exactly how PACAP promotes L-type channel opening.
We note that the mechanisms underlying intrinsic electrical activity are not yet resolved. Several reasonable explanations have been proposed based on the biophysical properties of channels known to be active during the inter-spike intervals, including voltage-gated Ca2+ channels (CaV1.3), BK channels, and Na+ channels (Lingle et al., 2018; Marcantoni et al., 2010; Milman et al., 2021). It is conceivable that PACAP, through a PLCε-dependent mechanism, harnesses one or more of these channels to increase the excitability of the chromaffin cell and the frequency of action potential spikes, thereby increasing intracellular Ca2+ and driving exocytosis.
Looming over the notion of intrinsic electrical activity is the question of its actual impact on the secretory response. A single action potential in a dissociated mouse chromaffin cell has been estimated to release between 1 and 2 granules (Moser and Neher, 1997); in adrenal slices, the number might approach 7 granules (Moser and Neher, 1997). One presumes that the increased levels of Ca2+ that accompany the transition from action potentials spikes to bursts would have a much greater impact on catecholamine release.
PACAP-dependent effects on the fusion pore
Peptide cargos are released more slowly into the extracellular space in cells stimulated by PACAP rather than ACh. However, the slowing effects of PACAP on fusion pore expansion must occur at a step that follows catecholamine release (Figs. 5 and 6). What mechanisms might be responsible for these observations? Elevations in cAMP production, of the sort presumably stimulated by PACAP, are associated with an increased incidence of transient fusion in rat pituitary lactotrophs (Calejo et al., 2013). Downstream of cAMP, there may be an Epac2-dependent recruitment of dynamin to the nascent fusion pore, as recently shown in pancreatic β cells (Gucek et al., 2019). Epac2 is expressed in the adrenal medulla (Fig. 8), and thus may play a similar role here as it does in the β cell. Dynamin has been previously reported by several groups, including ours, to act at fusion sites to slow both early and later stages of pore expansion (Anantharam et al., 2011; Fulop et al., 2008; Jackson et al., 2015; Tsuboi et al., 2004). Although a secretagogue-dependent recruitment of dynamin to fusion has not yet been demonstrated in chromaffin cells, our observations are consistent with such an outcome. Another possibility is that PACAP, by virtue of the small Ca2+ fluctuations it stimulates, preferentially triggers the fusion of vesicles bearing Syt7 (Anantharam and Kreutzberger, 2019). Such vesicles are notable for their increased sensitivity to Ca2+—thereby allowing fusion to occur when Ca2+ elevations are small—but also their propensity to discharge lumenal cargos slowly (Bendahmane et al., 2018; Rao et al., 2017; Tawfik et al., 2021).
PACAP stimulates a Gαs-coupled, PLCε-dependent pathway for Ca2+ influx and exocytosis
Alternative splicing of PAC1Rs impacts both ligand binding properties as well as the signaling cascade to which the receptor is coupled (Holighaus et al., 2011; Mustafa et al., 2007). However, the absolute requirement of PLCε here implies the participation of Gαs, rather than Gαq in pathway for chromaffin cell secretion (Oestreich et al., 2009; Schmidt et al., 2001; Zhang et al., 2011). Downstream of Gαs, our results also substantiate a role for the Rap/PLCε regulator, Epac (Smrcka et al., 2012). Pharmacological inhibition of Epac reduces PACAP-stimulated Ca2+ fluctuations, while pharmacological activation of Epac causes Ca2+ fluctuations that are qualitatively similar to those resulting from direct stimulation by PACAP. Thus, our results suggest that there must be an additional function for the Epac/Rap/PLCε signaling module in the chromaffin cell system which involves gating Ca2+ influx from the extracellular space (Fig. 4).
The absence of PLCε eliminates only the PACAP-stimulated component of the secretory response. However, this does not rule out a potential influence of PAC1 receptor activation on ACh receptor activity (Pugh et al., 2010; Starr and Margiotta, 2017). Our investigations were limited to short time window, which may be too brief for PACAP to exercise its function as regulator of gene transcription in the chromaffin cell (Carbone et al., 2019; Eiden et al., 2018). Longer term exposure of cells to PACAP or its signaling metabolites has also been shown to upregulate expression of machinery (e.g., Ca2+ channels) which might influence not only PACAP-stimulated exocytosis, but also ACh-stimulated exocytosis (Giancippoli et al., 2006; Novara et al., 2004). One should note that exactly how long PACAP released from splanchnic neurons persists at the synapse before it diffuses away during stress-activated synaptic activity is not a question that has been addressed, to our knowledge.
Summary
We have shown that two important neurotransmitters released from splanchnic neurons cause highly divergent Ca2+ responses and kinetically distinct fusion outcomes in chromaffin cells. These results are consistent with the evolving view of the adrenal medulla as producing nuanced and variable outputs, rather than stereotyped responses, to stressors that activate the sympathetic nervous system.
Acknowledgments
Christopher J. Lingle served as editor.
We thank Drs. Ronald W. Holz (University of Michigan) and Joseph Margiotta (University of Toledo) for critical comments.
Support was provided by National Institutes of Health grants R37DA0333987 to J.R. Traynor, R35GM127303 to A.V. Smrcka, R01GM111997/NS122534 to A. Anantharam, and American Heart Association grant 17GRNT33661156 to K.P.M. Currie.
The authors declare no competing financial interests.
Author contributions: A. Morales, R. Mohan, X. Chen, B.L. Coffman, L. Watch, J.L. West, J.R. Traynor, P.J. Kammermeier performed experiments. M. Bendahmane, S. Bakshi, J.R. Traynor, D.R. Giovannucci, K.P.M. Currie, and A.V. Smrcka devised experiments or helped implement experiments. D. Axelrod helped develop code for image analysis. A. Anantharam contributed to all aspects of the study and wrote the paper with feedback from A. Morales, R. Mohan, X. Chen, B.L. Coffman, M. Bendahmane, P.J. Kammermeier, D. Axelrod, K.P.M. Currie, and A.V. Smrcka.