Electrophysiological detection of nucleotide release from individual LDCVs. (A) Current (I)-voltage (V) relationship in a cell expressing P2X₂-EGFP. The current was activated by application of 0.2 mM ATP through a puffer pipette, and the I-V characteristics were then determined by ramping the membrane potential from -100 to +30 mV. The response obtained before application of ATP was subtracted from that recorded immediately after addition of the nucleotide to obtain the net current. The dotted horizontal and vertical lines represent the zero-current and the reversal potential, respectively. (B) Typical recording of current spikes evoked in a P2X₂-expressing cell by dialyzing the cell interior with a solution containing 2 μM free Ca²⁺. (C) Current spike obtained by averaging 32 recorded events (black line) with an amplitude ranging between 400 and 600 pA. (D) Cumulative number of current spikes recorded in four cells as shown in Fig. 1C (gray squares) and increase in whole-cell capacitance recorded in parallel (open circles) measured at 0.1 Hz. The scaling corresponds to 0.8 fF per spike (MacDonald et al., 2005).

Time course of nucleotide release. Exocytosis evoked by voltage-clamp depolarization in a P2X₂-EGFP-expressing Ins1-cell. (A) Membrane currents (I; middle) and increase in cell capacitance (ΔC) elicited by a 500 millisecond voltage-clamp depolarization from -70 mV to 0 mV (V; top). The inset shows the indicated part (highlighted by square) of the current trace after vertical and horizontal expansion. Asterisks indicate current spikes superimposed on the inward Ca²⁺-current (identified by eye). Spikes during the depolarization are smaller than at -70 mV due to the reduced inward driving force. The larger spikes cause brief artifacts on the capacitance trace that are unrelated to exo- or endocytosis. (B) As in A, but suramin (100 μM) was applied to the cell, resulting in nearly complete suppression of spikes in the current trace. (C) Cumulative histogram of the time (t) of the current spikes relative to the onset of the depolarization in eleven experiments as in A (circles). Overlaid are mono-exponential functions fitted to all points (time constant τ of 0.38 seconds; black line), and to the values greater than 500 milliseconds (τ of 0.28 seconds; dotted line). The shaded area indicates the duration of the depolarizing stimulus.

Time course of peptide release and luminal pH changes. (A,B) Cells expressing VAMP-pHluorin or IAPP-emerald (IAPP-EMD) were voltage-clamped and their footprint imaged at 10 Hz. Example images taken in separate cells transfected with IAPP-EMD (A) or VAMP-pHluorin (B) at the indicated times relative to the onset of a 500 millisecond depolarization from -70 mV to zero mV. (C) Time course of IAPP-EMD fluorescence (lower trace) in the ROI indicated by the circle in A in response to a 500 millisecond depolarization (top trace). Note that the IAPP-EMD fluorescence disappears after a delay of ≈2 seconds. (D) As in C for the experiment shown in B. Note that the VAMP-pHluorin trace rises abruptly to a plateau from which it declines mono-exponentially after a delay of ≈1 second. The gray line superimposed on the data points represents the best fit of a discontinuous function consisting of a straight-line segment followed by a mono-exponential decay to the remainder of the trace, using the first point of the rising phase as the start of the event. (E) Cumulative histograms showing the time, relative to stimulation, of the loss of IAPP-EMD fluorescence (squares), the increase in VAMP-pHluorin fluorescence (circles) analyzed as indicated by arrows in C and D. The functions superimposed on histograms in E are exponential functions fitted to the distributions yielding τ-values of 0.9±0.1 (circles), and 2.2±0.2 seconds (squares). For comparison, the function from Fig. 3B (fit to the nucleotide release data >500 milliseconds) is included as a dotted line.

Exocytosis does not necessarily lead to peptide release. (A,C) Simultaneous imaging of VAMP-pHluorin (top) and IAPP-ECFP (lower) in double-transfected cells. The cell (imaged at 10 Hz) was stimulated with a 3 second puff of solution containing 87 mM KCl. Times quoted below the images are relative to the peak in the VAMP-pHluorin signal. (A) Example of a granule (highlighted by a circle) that showed a transient increase in VAMP-pHluorin fluorescence and that culminated in the loss of IAPP-ECFP fluorescence. (B) Fluorescence intensities of IAPP-ECFP (open circles) and VAMP-pHluorin (black squares) within ROIs centered on the granule highlighted in A. (C) Example where the increase in VAMP-pHluorin fluorescence was not associated with the rapid loss of IAPP-ECFP fluorescence. (D) As in B, but for the granule shown in C. (E) Analysis of IAPP-ECFP fluorescence in 28 cells and in A,C. Data in the histogram were derived from traces as in B and D by subtracting the intensity measured at t=5.25 seconds from that at -1.0 second, both averaged over 0.5 second.

Parallel recording of nucleotide and peptide release from individual granules. (A,B) Confocal images of a section of the footprint of a voltage-clamped cell expressing both IAPP-pHluorin and P2X₂-mRFP. The images were recorded at the indicated times relative the onset of the IAPP-pHluorin flash. Exocytosis was elicited by intracellular dialysis of the cell with a buffer in which [Ca²⁺]_i was set at 2 μM. Note that the highlighted granules increase their fluorescence during the displayed sequence. In A, the signal from the granule is rapidly lost, while in B it remains elevated for several seconds. (C,D) Image sequences of a cell co-transfected with IAPP-pHluorin and P2X₂-mRFP1 and stimulated as in A and B. The entire cell was imaged in the IAPP-pHluorin channel (see Materials and Methods), and whole-cell current spikes due to activation of the P2X₂ receptors were recorded in parallel. Examples of a short-lived fluorescence transient in C, and a long-lasting event in D. (E,F) Time course of the average fluorescence intensity in the ROIs indicated by the white circle in C and D (top) and inward membrane currents associated with the events (lower). Note slow decay of fluorescence in F. (G) Histogram of the decay constants of 56 events as in C and D. The shaded area indicates decay constants greater than 350 milliseconds. (H) Cross-correlation histogram of the times of fluorescence peaks vs the times of current peaks in three experiments similar to that shown in C and D.