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Calcium microdomains generated by tight clusters of calcium channels regulate fusion of small vesicles at the synaptic terminal and have also been suggested to trigger exocytosis of large dense-core vesicles from neuroendocrine cells. To test this idea, we have compared sites of exocytosis and the spatial distribution of calcium channels in chromaffin cells. Fusion of individual vesicles was visualized using interference reflection microscopy and the submembranous calcium signal was assessed using total internal reflection fluorescence microscopy. Depolarization triggered a burst of exocytosis from up to seven sites in a membrane area of 11 μm 2, but these sites did not colocalize with calcium microdomains. Instead, calcium influx occurred in large patches (averaging 34 μm 2) containing a mixture of P/Q- and N-type channels. About 20% of fusion events occurred outside calcium channel patches. Further, the delay between the onset of stimulation and a burst of exocytosis was prolonged for several seconds by increasing the concentration of the slow calcium chelator EGTA from 1.5 to 5 m m.

These results demonstrate that while calcium channels and release sites tend to congregate in specialized regions of the surface membrane, these have dimensions of several micrometres. The dominant calcium signal regulating release in chromaffin cells is generated by the cooperative action of many channels operating over distances of many micrometres rather than discrete clusters of calcium channels generating localized microdomains. Introduction It is unclear how secretion in neuroendocrine cells is coupled to the opening of Ca 2+ channels. One view is that a docked vesicle is triggered to fuse by a nearby ‘Ca 2+ microdomain’ generated around a tight cluster of open Ca 2+ channels , leading to the suggestion that excitation–secretion coupling in neuroendocrine cells is similar to synapses, where vesicles and Ca 2+ channels cluster tightly at active zones (;; ). However, this view is difficult to square with the functional and morphological differences in excitation–secretion coupling at synapses and endocrine cells.

At synapses, vesicles begin fusing. Cell preparation and solutions Bovine adrenal chromaffin cells were prepared as described. Adrenal glands were obtained from a local slaughterhouse and chromaffin cells isolated by collagenase digestion followed by purification using a Percoll gradient. Cells were plated onto poly- l-lysine-coated coverslips and used 1–3 days after preparation. Ringer solution contained (in m m): 140 NaCl, 2.5 KCl, 1 MgCl 2, 10 Hepes, 10 glucose, 2.5 CaCl 2, pH 7.4, 300 mosmol kg −1.

For TIRFM experiments 1 n m tetrodotoxin (Tocris, Ellisville, MO, USA) was added to Ringer solution. Nifedipine was from Sigma (St Louis, MO, USA), nimodipine from EMD Biosciences (San Diego, CA, USA); ω-agatoxin IVA and ω-conotoxin GVIA were from Alomone Labs (Jerusalem, Israel). Solutions were perfused onto cells using an array of quartz pipes (World Precision Instruments, Sarasota, FL, USA) controlled by a system of valves (Warner Instrument Corp., Hamden, CT, USA). The solution in the patch pipette contained (in m m): 120 caesium methanesulfonate, 10 TEA-Cl, 5 MgCl 2, 20 Hepes, 3 Na 2ATP, and 1 NaGTP, pH 7.2, 290 mosmol kg −1. For IRM experiments, the pipette solution contained variable amounts of Ca 2+ buffer, either (in m m) 0.1 EGTA, 1.5 EGTA, 5 EGTA or 0.4 BAPTA. For TIRFM experiments, pipette solution contained either 0.1 m m OregonGreen488-BAPTA-2 ( K d= 0.6 μ m; OGBAPTA-2) or 0.2 m m Fluo-5N ( K d= 90 μ m; Molecular Probes, Eugene, OR, USA). Electrophysiology and imaging IRM imaging was performed as described previously (; ).

Cells were voltage clamped in the whole-cell configuration using an Axopatch 200A amplifier (Axon Instruments, Union City, CA, USA) and acquired with a G4 Macintosh equipped with an ITC-16 interface (Instrutech Corp., Port Washington, NY, USA) controlled by the Pulse Control extension of IGOR Pro software (Wavemetrics, Lake Oswego, OR, USA). Exocytosis was elicited by depolarizing from −80 mV to +10 mV, either as a single 500 ms step or as a train of 24 × 105 ms or 4 × 90 ms pulses. We used 105 ms pulses to allow each to be delivered at the beginning of a frame acquired by the camera. Image acquisition was performed using IPLab software and triggered by a TTL pulse delivered by the ITC-16 at the beginning of the electrophysiological recording.

Calcium currents were leak subtracted using macros written in IGOR Pro. All IRM images and movies were ‘background subtracted’ using a macro written in IPLab (Scanalytics, Fairfax, VA, USA) to better visualize changes. ‘Through the objective’ TIRFM was combined with IRM as described (; ).

A 488 nm beam from an argon laser provided the TIRFM excitation light. The light for IRM originated from a 100 W xenon lamp (Newport-Oriel, Irvine, CA, USA) and was transmitted through a 510WB40 filter (Omega Optical, Brattleboro, VT, USA) before passing through a combining cube (Oriel). The filter block contained a 505DRLP dichroic (Omega), which reflected 95% at 488 nm and about 35% at 510 nm, while transmitting 80% at wavelengths 515 nm.

About 35% reflection was sufficient for IRM if the xenon lamp was at maximum power. When only TIRFM was used, emitted light was filtered through a HQ510LP filter (Chroma Technology, Rockingham, VT, USA) and 488 nm notch filter (Coherent, Santa Clara, CA, USA). When TIRFM was combined with IRM, emitted light was filtered through a 545AF75 filter (Omega).

Images were magnified × 2.2 before being acquired by a Princeton Instruments Pentamax CCD camera and Winview32 acquisition software (Roper Scientific, Trenton, NJ, USA). Image acquisition was synchronized with the electrophysiological recording using a Master-8 pulse generator (AMPI, Jerusalem, Israel).

The membrane current and camera exposure signal time were digitized (20 kHz) and filtered (5 kHz) by a Digidata 1322A interface (Axon). Images of Ca 2+ influx in – are shown as Δ F/ F 0, where Δ F= F– F 0, F is the individual frame and F 0 is the average image at rest (obtained from between 4 and 15 consecutive frames acquired before the stimulus). The protocol for identifying sites of calcium influx by TIRFM is described further in Supplemental. To quantify changes in Ca 2+ influx between different stimulus trains ( and ), we compared Δ F values measured in the frame in which the 2 ms stimulus was applied. Each pixel from the Δ F image obtained after adding a blocker of Ca 2+ channels was divided by the corresponding pixel in the Δ F image obtained before addition of the blocker. The ratio values for each pixel within the Ca 2+ channel patch were then averaged to give the mean intensity ratio (± s.d.) over all pixels in the patch.

The ratio image of Stim 2/Stim 1 therefore provided a measure of the relative change in the amount of Ca 2+ influx in the footprint. Measurements of the free Ca 2+ under the membrane using Fluo-5N were calibrated by measuring F min in pipette solution containing 10 m m EGTA and F max in pipette solution containing 1 m m CaCl 2.

The K d for Fluo-5N was assumed to be 90 μ m, measured by the manufacturers. Results are expressed as mean ± s.e.m. Unless otherwise indicated. Absence of L-type calcium channels in the footprint A, images of Ca 2+ influx (Δ F/ F) obtained at rest, then after successively adding 20 μ m nifedipine and 20 μ m nimodipine ( C), 1 μ mω-conotoxin GVIA ( E) and 200–400 n mω-agatoxin IVA ( G), all to the same cell. The corresponding ratio images are shown in D, F and H; the Ca 2+ currents are shown in B.

Dihydropyridines blocked the majority of Ca 2+ channels in the surface membrane of this cell as a whole ( B), but none in the footprint ( D). Calibration bar in B: 200 pA × 2 ms; scale bar, 5 μm; ratio calibration bar = 1.5 (white), 1.0, 0.5, 0 (black). Calcium channel patches in footprints consist primarily of N- and P/Q-type channels A– D, images of Ca 2+ influx (Δ F/ F) obtained at rest (Stim 1, top row) and in a test condition (Stim 2, second row), as well as the ratio of Stim 2/Stim 1 (Ratio, third row). Each frame is averaged from 6 responses to depolarizations lasting 2 ms.

Before and during the second stimulus set (Stim 2), cells were perfused either with normal Ringer solution ( A), 20 μ m nifedipine and 20 μ m nimodipine ( B), 1 μ mω-conotoxin GVIA ( C) or 200–400 n mω-agatoxin IVA ( D). Mean ratio values (± s.d.) over the patch of Ca 2+ channels are shown below the ratio images. The averaged whole-cell Ca 2+ currents for Stim 1 (black) and Stim 2 (red) are at bottom; all calibration bars: 200 pA × 2 ms. Scale bar: 5 μm; ratio calibration bar = 1.5 (white), 1.0, 0.5, 0 (black).

E, collected results ( n= 28 cells), showing the percentage change in Ca 2+ influx over the whole cell (Δ Q Ca) versus the intensity ratio over the patch of Ca 2+ channels. Small crosses represent data from individual cells perfused with either control Ringer solution (black), dihydropyridines (red), conotoxin (blue), or agatoxin (green) before collecting the second set of stimulus images. Large markers with error bars represent means. The diagonal line marks the relationship expected if Δ F/ F were directly proportional to Q Ca and all types of Ca 2+ channels were randomly distributed over the whole surface of the cell, including the footprint. Visualizing sites of calcium influx by TIRFM A, images showing the relative change in fluorescence (Δ F/ F) of OGBAPTA-2 (0.1 m m) in the footprint of a chromaffin cell. Averaged from six movies of 7 × 11 ms frames acquired at 10 s intervals, all from the same cell. Calcium influx was triggered by a 2 ms depolarization (−80 to +10mV) delivered at the beginning of frame 5; a 1 ms prepulse from −80 mV to +150 mV was delivered immediately preceding the 2 ms depolarization.

Red lines indicate the edges of the footprint. B, whole-cell Ca 2+ currents recorded for each of the 6 stimuli delivered to the cell in A (black traces) and the average of the 6 traces (red). C, five individual examples of the stimulus frames used to make the averaged frame 5 in A. D, a gallery of averaged stimulus frames showing the relative change in intensity (Δ I/ I 0) from 10 different cells on a grey scale. Each image is the average of 6 or 8 stimulus frames.

Scale bar in arbitrary units shows Δ I/ I 0. Specialized sites of fusion A and B, background-subtracted images from the IRM movies of two cells, each stimulated twice (Stim 1 and Stim 2). Arrows are located in the same position during the first and second stimulations, indicating fusion events occurring in the same spot. Stim 1 and Stim 2 images in B are from Supplemental (available online only). C, histogram showing the number of fusion events that occurred within the same spots two, three, or four times, from a total of 307 events. The bar marked ‘1×’ is the number of events that did not overlap with any others.

Using IRM to detect fusion of large dense-core vesicles To investigate excitation–secretion coupling in chromaffin cells we visualized sites of fusion using IRM, a technique that detects local invaginations of the surface membrane generated by fusion of a vesicle. Examples of such events are shown in, where they appear as localized ‘spots’ within the footprint of the cell (see in Supplemental material, available online only). A large number of experimental observations indicate that these spots are generated by the Ω-shapes formed when a vesicle fuses with the surface membrane (, ). First, spots in the footprint only occur during and after the application of a depolarizing stimulus.

Second, they are synchronized in time and space with the release of FM4-64, acridine orange or pro-atrial natriuretic peptide (ANP)-GFP within granules, as visualized by simultaneous TIRFM and IRM imaging. Third, the number of events in a footprint is directly proportional to the total increase in surface area measured by the capacitance technique and follows the same time course. Fourth, the average capacitance increase per event is 1.43 fF, which is very similar to the average capacitance of a granule estimated. Fifth, the kinetics of IRM signals are altered by a number of experimental manipulations known to affect the fusion or retrieval of granules. For instance, the speed of IRM signals are calcium dependent: recovery of the signal is blocked by strontium, which also blocks vesicle retrieval assayed by capacitance. The full procedure by which IRM was used to identify the timing and location of fused granules is described in the online ‘’ and Supplemental.

Bursts of fusion events occurring in clusters on the membrane A and B, background-subtracted IRM images from 2 different cells stimulated with a train of 24 × 105 ms depolarizations at 5 Hz. Before stimulation, IRM footprints were uniformly dark (top row), but discrete spots appear on depolarization.

A was recorded with 1.5 m m EGTA in the patch pipette and B with 5 m m EGTA. Image in B comes from Supplemental, available online only.

Scale bar, 5 μm. C and D, the cumulative number of fusion events and the corresponding whole-cell Ca 2+ currents plotted for cells in A and B. Coloured arrows correspond to time points at which the frames shown in A and B were extracted. Note the slower inactivation of Ca 2+ currents in D compared to C. Scale bar, 5 fusion events. Bursts of exocytosis were delayed by EGTA When a chromaffin cell was depolarized, fusion events within the footprint often occurred in bursts, and the timing of these depended on the concentration of the Ca 2+ chelator inside the whole-cell pipette.

The cell in was dialysed with 1.5 m m EGTA, which binds Ca 2+ ions with a K d of ∼150 n m. While Ca 2+ channels were closed, there was no activity (top), but a stimulus train lasting 5 s triggered the appearance of discrete bright spots. A cumulative plot of fusion events shows that exocytosis in this footprint occurred in two distinct bursts, the first complete within 1 s of the beginning of the stimulus train and the second occurring more slowly over a period of several seconds. Two phases of exocytosis are also observed in chromaffin cells using the capacitance technique and release of caged Ca 2+: the initial exocytic burst is complete within a few hundred milliseconds at 10–20 μ m Ca 2+ and the second occurs at a rate of 0.1 s −1. In comparison, we estimated that the Ca 2+ within ∼100 nm of the membrane reached ∼20–30 μ m during depolarization under conditions of low Ca 2+ buffering. Visualizing calcium influx with a low affinity calcium indicator A, ten consecutive frames showing Δ F/ F from a cell loaded with 0.2 m m Fluo-5N through the patch pipette.

The whole-cell Ca 2+ current is shown below. The four 90 ms depolarization steps occurred within frames 29, 31, 33 and 35 (100 ms per frame). Scale bar: 5 μm. B, the submembranous Ca 2+ averaged over the whole footprint (black) for the same cell shown in A was compared with the Ca 2+ of two ROIs (each 10 pixels × 10 pixels), one ROI centred over a patch of Ca 2+ channels (green box in A) and one remote from Ca 2+ channels (red box). The black bar indicates the time of the depolarization. Similar results were seen for seven other cells. The cumulative exocytosis events measured by IRM in response to the same stimulus is shown above ( n= 10 cells).

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When the concentration of EGTA inside the pipette was increased from 1.5 m m to 5 m m, the initial rate of exocytosis was slowed. Averaged time courses of exocytosis in 0.1 m m, 1.5 m m and 5 m m EGTA are shown by the cumulative histograms in. Each plot is averaged from 4–5 footprints, and was obtained by making cumulative histograms of all events under each condition, then dividing by the number of footprints from which these measurements were collected. The initial rates of granule fusion were then compared by making linear fits to the cumulative histograms (bold lines in ). Under these stimulation conditions, the frequency of fusion events was sensitive to EGTA, as would be expected from capacitance measurements. The initial rate of fusion in 0.1 m m EGTA was sevenfold higher than in 5 m m EGTA , but there was still an obvious burst of exocytosis in 5 m m EGTA; the crucial difference was that the most rapid phase of fusion in 5 m m EGTA was delayed by 3–4 s. In other words, the slowing of exocytosis by EGTA could be overcome by more prolonged Ca 2+ influx.

This observation argues strongly against the idea that the signal triggering granule fusion is a localized microdomain, for two reasons. First, both theory and experimental measurements (; ) demonstrate that Ca 2+ microdomains persist in EGTA concentrations up to 5 m m, because this chelator binds Ca 2+ ions relatively slowly. The major effect of EGTA is to suppress the ‘global’ increase in Ca 2+ generated by diffusion of Ca 2+ away from sites of influx (; ).

Second, Ca 2+ microdomains are generated within milliseconds when Ca 2+ channels open, even in the presence of high concentrations of EGTA. The delayed burst of exocytosis in 5 m m EGTA must therefore reflect the gradual accumulation of free Ca 2+ ions some distance from calcium channels. EGTA increased the delay between the beginning of stimulation and the first burst of exocytosis A– C, cumulative plot of number of fusion events per footprint observed during trains of 24 × 105 ms depolarizations delivered at 5 Hz. Cells were dialysed with 0.1 m m EGTA ( A; n= 4), 1.5 m m EGTA ( B; n= 5), 5 m m EGTA ( C; n= 4) or 0.4 m m BAPTA ( A; n= 3). D, the traces in A–C superimposed on an expanded time scale.

The initial phases of release have been fitted with straight lines through the origin (bold). E, the initial rate of fusion events (per second per footprint) as a function of the concentration of EGTA or BAPTA in the patch pipette. The line through the points is the least-squares fit of the relation R= R max1 − ( E/( E+ E 1/2)), where E is the concentration of buffer in m m, E 1/2 is the concentration where inhibition is half-maximal (0.36 m m) and R max is the maximum rate of release in the absence of any calcium buffer in the patch pipette (16 events per second per footprint). Both R max and E 1/2 were free parameters in the fitting process. The red point is a measurement in BAPTA and the black points are measurements in EGTA. Error bars show the standard deviation of the linear fits in D (some of these are narrower than the points).

Secretion triggered by calcium microdomains at the synapse can be inhibited much more efficiently by the Ca 2+ chelator BAPTA compared to EGTA (; ). BAPTA binds Ca 2+ ions with a similar affinity to EGTA but about 60-fold faster, and therefore has a stronger effect on calcium signals close to calcium channels. To compare the effects of these buffers on granule fusion in chromaffin cells, we first looked at the relation between the initial rate of fusion and the EGTA concentration, shown by the black circles in. These measurements could be described empirically by a function assuming that EGTA inhibits granule fusion according to the relation R= R max1 − ( E/( E+ E 1/2)), where E is the concentration of EGTA in m m, E 1/2 is the concentration where inhibition is half-maximal (0.36 m m) and R max is 16 events per second per footprint. To test whether BAPTA was any more effective than EGTA at slowing granule fusion, we introduced 0.4 m m BAPTA, which would be expected to have an effect that was approximately half-maximal if the two chelators were equally effective at antagonizing granule fusion.

The red point in shows that BAPTA was not significantly more, or less, effective than EGTA. This observation indicates that the equilibrium dissociation constant of these chelators is more important than the association constant for Ca 2+ binding in inhibiting granule fusion, and that this process is therefore triggered by the global increase in Ca 2+ rather than Ca 2+ microdomains. Repeated fusion events at specialized sites At some sites in the membrane, fusion events occurred repeatedly during a train of stimuli lasting 5 s.

Shows examples of two cells which were stimulated twice, and the arrows identify fusion events which occurred in the same spot in the separate stimuli (see also online Supplemental ). Of 307 fusion events in 12 cells stimulated 3–6 times, 115 occurred within one pixel of each other (the centre of the event being defined as the pixel in which the IRM signal was at its maximum and each pixel being 73 nm square).

In some cases, different fusion events occurred centred over the same pixel three or more times. Thus at least 37% of fusion events occurred at sites that were fixed in the membrane over the tens of minutes time scale of these experiments. This figure will be a lower limit because the relatively low average release probability at a single site would have required many repetitive stimuli to identify all the preferred sites in this way. Nonetheless, these results indicate that the membrane of chromaffin cells contains areas at which vesicles can fuse repeatedly.

Clustering of multiple fusions in a single site could be due to the sequential arrival of different granules to preferential release zones of ∼0.4 μm 2 (; ) and/or to a small granule population undergoing repeated cycles of fusion and re-sealing in a circumscribed portion of the membrane. Calcium influx occurred in large patches We next sought to investigate how sites of fusion were organized spatially relative to Ca 2+ channels in the membrane, and in particular whether depolarization generated discrete calcium microdomains or more diffuse signals. Submembranous Ca 2+ was imaged by TIRFM using the indicator OGBAPTA-2 (0.1 m m).

At the synapse of goldfish retinal bipolar cells, Ca 2+ microdomains generated by tight clusters of Ca 2+ channels can be clearly visualized after depolarizations lasting 20 ms (; ). We sought to capture an even briefer ‘snap-shot’ of Ca 2+ entry in chromaffin cells by depolarizing for just 2 ms and integrating the signal over a single frame lasting 11 ms. This stimulus moved a Ca 2+ charge of 0.9 ± 0.1 pC into the cell, similar to the ∼1 pC delivered in response to a single action potential.

The resulting calcium signal is shown in, which is a series of 5 out of 7 consecutive frames, with the depolarization delivered at the beginning of frame 5. Calcium influx was not uniform across the footprint, but occurred in distinct patches. Then, after Ca 2+ channels closed, the fluorescence gradually dissipated as Ca 2+ diffused away from entry sites (, frames 6 and 7). The source of Ca 2+ influx was a relatively fixed array of channels because when the depolarization was repeated , the submembranous Ca 2+ signal was similar for each of five stimuli. Calcium influx therefore occurred diffusely rather than in discrete puncta.

Calcium influx visualized by TIRFM is also shown in Supplemental. Might these ‘patchy’ Ca 2+ signals reflect non-uniform attachment of the footprint to the glass coverslip? Three observations argue against this possibility. First, the IRM experiments showed that footprints presented a homogeneous pattern of interference before stimulation. Moreover, signals arising exclusively from the edges of the footprint were not considered because these were the membrane regions that tended to display a looser attachment to the coverslip. Second, the fluorescence intensity of OGBAPTA-2 viewed by TIRF was relatively uniform across the footprint at resting levels of Ca 2+ ( below).

And, third, the submembranous Ca 2+ signal measured by TIRFM was also relatively uniform when a large Ca 2+ load was introduced and allowed to dissipate after closure of Ca 2+ channels. An example of this behaviour can be seen in, where there is a patchy Ca 2+ signal evident during the first depolarization of a train (frame 29), but a much more uniform signal after closure of Ca 2+ channels at the end of the train (frame 36). We conclude that the patchy signals reported by OGBAPTA-2 reflected the distribution of calcium channels in the surface membrane. Defining the area of the footprint and calcium patches A, defining the area of the footprint. TIRFM image of a voltage-clamped chromaffin cell, at rest, loaded with 0.1 m m OGBAPTA-2. To acquire TIRFM images at 11 ms per frame, data collection was restricted to a 50 pixels × 50 pixels ROI (green box, left). To define the outline of the footprint, we determined the minimum (MIN frame1) and maximum (MAX frame1) pixel intensities for the first frame in a movie.

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A threshold was then set at the minimum pixel intensity (MIN frame1) + 20% of the total intensity range (MAX frame1) – (MIN frame1). All pixels below the threshold are marked in red (middle), and the margins of these regions are shown by the lines in image at right. B, defining the area of calcium influx. An image of the relative change in the fluorescence of OGBAPTA-2 (Δ F/ F) in the frame in which Ca 2+ influx was activated for 2 ms is shown at left. This image was thresholded by first determining the maximum pixel intensity over all the prestimulus frames (MAX frame1-4) and the maximum pixel intensity of the stimulus frame (MAX frame5). The minimum threshold was set as (MAX frame1-4) + 20% of the intensity range (MAX frame5) – (MAX frame1-4). The pixels above this threshold are marked in green in the middle panel and the margins of this area are shown by the green line in the right panel.

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Measuring areas of calcium influx The area covered by arrays of Ca 2+ channels varied widely. Two relatively localized Ca 2+ signals, indicating a small cluster of Ca 2+ channels, are shown by the red arrows in. These signals might be termed ‘microdomains’ because their spatial scale is 1–2 μm , but the predominant signal was much more widely distributed. The areas of Ca 2+ influx in the footprints of ten other cells are shown in. Surveying these examples, it is possible to see ‘hotspots’ within the larger regions of Ca 2+ influx, indicating that the density of Ca 2+ channels per unit area was not necessarily uniform across these patches.

The basic observation was that independent microdomains (such as arrowed in ) were rare, and the predominant Ca 2+ signal was distributed over much wider regions. These patches of Ca 2+ influx were relatively well circumscribed and their boundaries could be recognized by eye. To measure the areas of the footprint and patches of Ca 2+ influx we applied a thresholding technique. Shows the resting fluorescence of a chromaffin cell loaded with 0.1 m m OGBAPTA-2 (left).

To allow imaging at 11 ms per frame, we sampled a 50 pixels × 50 pixels region of interest (ROI). The minimum and maximum pixel intensities for the first frame in a movie were measured (MIN frame1 and MAX frame1), and a threshold set at 20% of the total intensity range (i.e. MIN frame1+ 0.2(MAX frame1− MIN frame1)). This thresholding criterion was chosen as one which effectively separated the footprint from the background signal over the neighbouring coverslip. An example is shown in, where all pixels below the threshold are marked in red (middle), and the margins of these regions, marking the outline of the footprint, are shown by the lines in the image on the right. Similar criteria were applied to define patches of Ca 2+ influx. Shows the relative change in the fluorescence of OGBAPTA-2 in the frame in which Ca 2+ influx was activated for 2 ms (left).

First we determined the maximum pixel intensity over all the prestimulus frames (MAX frame1-4), then the maximum pixel intensity of the stimulus frame (MAX frame5). The minimum threshold was set at 20% of this range (i.e. MAX frame1-4+ 0.2(MAX frame5− MAX frame1-4)). The pixels above this threshold are marked in green in the middle panel in, and the margins of the contiguous area are shown by the green line in the right panel.

The size of the Ca 2+ channel arrays ranged from 10 to 78 μm 2, with a mean size of ∼34 μm 2. On average there were two distinct patches per footprint. Exocytosis within patches of calcium channels The results presented in, and indicated that clusters of fusion events occurred on spatial scales similar to patches of Ca 2+ channels. To determine more precisely where exocytosis occurred relative to Ca 2+ channels, we applied IRM and TIRFM in the same cells.

Three examples are shown in. First, the IRM images at the top were obtained by integrating over a period of 12 s from the onset of the stimulus, which was delivered in the perforated patch configuration.

The longer integration time compared to provided a view of all the sites where granules fused over this period, and the brighter white spots reflect sites where two or more fusion events occurred (as assessed by observing the movie of difference images frame by frame, as shown in ). The corresponding TIRFM images at the bottom show Ca 2+ channel patches visualized in the same footprint by going whole-cell with the pipette containing 0.1 m m OGBAPTA-2. Overlaid onto the Ca 2+ channel patches are yellow and blue spots marking the sites of fusion events occurring during the stimulus (synchronous) and after the stimulus (asynchronous), identified from the IRM movies. Most sites of fusion were within the large regions of the surface membrane containing Ca 2+ channels. Fusion events occurred preferentially close to calcium channels IRM images averaged over a 12 s period from the start of the stimulus are shown in the top row and areas of Ca 2+ influx visualized by TIRFM are shown below for three different cells ( A– C). To trigger the fusion events ( A–C, top), cells were voltage clamped in the perforated-patch configuration and stimulated with 4 × 90 ms depolarizations at 5 Hz.

We then broke-through to whole-cell configuration and loaded cells with 0.1 m m OGBAPTA-2. The TIRFM images of Ca 2+ influx ( A–C, bottom) show the average stimulus frames (Δ F/ F), as in. Overlaid onto the TIRFM images are the sites of synchronous (yellow) and asynchronous (blue) fusion events visualized by IRM, the outline of the footprints (red lines), and the perimeter of the Ca 2+ channel patches (green lines). D, average number, per cell, of asynchronous and synchronous fusion events occurring inside vs. Outside of calcium channel patches. The differences between the means for asynchronous events inside vs. Outside were significantly different ( t test, P= 0.006), as were the differences between the means for synchronous events inside vs.

Outside ( P= 0.03). Error bars are s.e.m.; n= 10 cells for each. E, total number of fusion events recorded for 10 cells (137 total events) is broken down to reveal the ratio of synchronous to asynchronous, inside and outside of the calcium channel patches. To determine whether fusion events occurred preferentially within the Ca 2+ channel patches, we measured the area of the footprint ( A foot) and the area covered by patches of Ca 2+ channels ( A patch) for each cell ( n= 10). If all of the fusion events visualized under IRM ( N total= 137 events) were randomly distributed across the footprint, then the predicted number of fusion events within A patch would be: N predict= ( A patch/ A foot) × N total. The actual number of fusion events occurring within the Ca 2+ channel patch ( N actual) was compared to N predict for each cell.

The total N actual was 109 events, significantly larger ( P= 0.01, Student's paired, two-tailed t test) than the total N predict of 92 events. In 70% of the cells, N actual N predict, while in the remaining 30% N actual= N predict. These results confirmed that fusion events in the footprints of chromaffin cells were not randomly distributed but occurred preferentially within patches of Ca 2+ channels. Might the clustering of fusion events within Ca 2+ channel patches simply reflect the higher local Ca 2+ concentration? Two observations argued against this possibility. First, while most fusion sites occurred in regions of the surface membrane containing Ca 2+ channels, several were towards the margins of these patches (e.g.

) and 20% outside (e.g. Second, when we analysed all the fusion events in 10 cells, we found that asynchronous events also occurred preferentially within patches of Ca 2+ channels. These results indicate that the local Ca 2+ concentration was not the only factor causing fusion to occur preferentially in regions of the surface membrane containing Ca 2+ channels. It seems most likely that greater numbers of vesicles were docked in these areas or that the vesicles were better primed for release. The calcium signal triggering synchronous and asynchronous fusion events The stimulus we used to visualize the distribution of Ca 2+ channels – a depolarization lasting 2 ms – was too brief to trigger exocytosis. To better understand excitation–secretion coupling we therefore also imaged the submembranous Ca 2+ signal during more prolonged Ca 2+ influx sufficient to trigger exocytosis, using trains of 4 × 90 ms depolarizations. Fluo-5N was used as the Ca 2+ indicator because its relatively low affinity ( K d= 90 μ m) prevented saturation in the face of these much larger Ca 2+ loads.

Shows a series of consecutive frames lasting 100 ms, with 90 ms depolarizations delivered during frames 29, 31, 33 and 35. Patches of Ca 2+ channels could be recognized in response to the first stimulus in the train (frame 29), but diffusion of Ca 2+ from these regions caused a gradual build-up under the whole membrane (compare frames 30 and 36). Shows the submembranous Ca 2+ concentration over the whole footprint (black) compared with two 10 pixels × 10 pixels ROIs, one within (green) and one outside (red) a patch of Ca 2+ channels. Average Ca 2+ levels reached a maximum of about 30 μ m within the Ca 2+ patch during the stimulus, but rapidly fell after closure of Ca 2+ channels.

Of a total of 137 fusion events, 43% occurred 1 s or more after Ca 2+ influx had stopped, at Ca 2+ concentrations below 6 μ m. Clearly, exocytosis was not tightly coupled to the opening of Ca 2+ channels.

These results provide further evidence against the idea that fusion of secretory vesicles in chromaffin cells is driven by localized Ca 2+ microdomains. Calcium channel patches consisted primarily of N- and P/Q-type channels Bovine adrenal chromaffin cells express three types of voltage-activated Ca 2+ channels, N-, L- and P/Q-type. Several studies have examined the efficiency with which the different channel subtypes trigger exocytosis in chromaffin cells but with conflicting results (;; ). N- and P/Q-type channels contain a synprint region that allows interaction with syntaxin, SNAP-25, and synaptotagmin ; thus, these channels may localize to regions of the surface membrane where the SNARE complex has formed in preparation for fusion. We therefore compared how different subtypes of Ca 2+ channels were distributed within and between patches.

After imaging sites of Ca 2+ influx under normal conditions, we applied dihydropyridines (DHP) to block L-type channels, ω-conotoxin GVIA (CTX) to block N-type channels, or ω-agatoxin IVA (AGA) to block P/Q-type channels. The averaged responses to the first set of 2 ms stimuli under control conditions are shown in the top row in (‘Stim 1’); the response to the second set of stimuli under test conditions is shown in the middle row (‘Stim 2’), and the ratio images of Stim 2/Stim 1 are at the bottom. When the second set of stimuli was also delivered in the absence of any pharmacological agents, the average ratio of (Stim 2/Stim 1) was one , indicating there was little change in Ca 2+ influx between the two sets of stimuli (, bottom). The (Stim 2/Stim 1) ratio was also equal to one after treatment with dihydropyridines, indicating that blocking L-type Ca 2+ channels did not significantly decrease Ca 2+ influx in the footprint, even though the whole-cell Ca 2+ current was inhibited by 20%. In contrast, treatment with CTX resulted in an 80% decrease in Ca 2+ influx between Stim 1 and Stim 2 , while application of AGA led to a 40% decrease. These results indicate that patches of Ca 2+ channels within the footprint were made up primarily of N- and P/Q-types, with very few active L-type channels.

The ratio images (Stim 2/Stim 1) in were relatively uniform, as they were in 13 other cells tested, indicating that the distributions of N- and P/Q-type channels were similar across a patch. Compares how these blockers of Ca 2+ channels affected the total amount of Ca 2+ influx into the cell (measured as the integral of the Ca 2+ current, Q) and the influx into the footprint (measured as the mean pixel intensity in the ratio image Stim 2/Stim 1). In cells treated with CTX and AGA, inhibition of the Ca 2+ current was correlated with a decrease in the amount of Ca 2+ entering through the footprint.

In contrast, in cells treated with DHP, Ca 2+ influx into the footprint was not significantly reduced. Thus, L-type channels were segregated away from N- and P/Q-type channels in the footprint. This conclusion was reinforced by successively blocking each Ca 2+ channel subtype in the same cell. In the example in, applying DHP reduced the total Ca 2+ current by about 70% , without any effect on the Ca 2+ influx through the footprint (mean pixel intensity ratio of 0.99 ± 0.13 in ). When CTX was added in addition to DHP, the mean intensity ratio was 0.59 ± 0.12 , while further addition of AGA completely blocked Ca 2+ influx. Thus the footprint contained only N- and P/Q-type channels, and the ratio images show that these were distributed relatively uniformly.

Discussion The results of this study demonstrate that the time and place of vesicle fusion in chromaffin cells is not tightly coupled to the opening of Ca 2+ channels in the surface membrane. Direct visualization of the submembranous Ca 2+ signal associated with brief openings of Ca 2+ channels demonstrated that these were not discrete microdomains, but large areas, often covering tens of square micrometres. Three aspects of the results obtained with Ca 2+ buffers also argued against tight coupling between sites of granule fusion and Ca 2+ channels within these large regions of the membrane.

First, vesicle fusion in chromaffin cells was very sensitive to EGTA , a chelator that binds Ca 2+ ions relatively slowly and which is therefore only effective at reducing the Ca 2+ signal at micrometre distances from Ca 2+ channels. Second, the rate of vesicle fusion accelerated as Ca 2+ accumulated in the cytoplasm over periods of seconds. The time scale over which calcium microdomains are generated is three orders of magnitude shorter. Third, BAPTA, a fast chelator of Ca 2+ ions, was no more effective at slowing the rate of exocytosis than EGTA, which binds Ca 2+ ions about 60-fold more slowly. Although the majority of fusion sites were located within areas of the surface membrane containing patches of Ca 2+ channels, exocytosis also occurred at sites remote from channels. In addition, asynchronous release after closure of Ca 2+ channels was an obvious feature of exocytic responses , providing direct evidence that release did not require Ca 2+ microdomains generated by closely coupled Ca 2+ channels.

All these observations run counter to the suggestion that fusion of large vesicles in chromaffin cells is driven by localized Ca 2+ signals similar to those observed at the synapse (; ). The results we have presented stand in contrast to those of, who reported that brief (. Calcium channels distributed over large areas of membrane On depolarization, calcium influx occurred through areas of the surface membrane covering many tens of squared micrometres, and ‘microdomains’ could not be detected. TIRFM has reliably detected calcium microdomains at the active zones of retinal bipolar cells and hair cells (; ), so the lack of such signals in chromaffin cells is unlikely to be a limitation of the technique. Although imaging indicated that the density of channels within a patch was not uniform ( and ), it was clear that the Ca 2+ signal over most areas of the membrane was affected by Ca 2+ influx over many channels distributed widely.

How many channels formed a patch in a chromaffin cell? The average surface area of a chromaffin cell is about 710 μm 2 (assuming a specific membrane capacitance of 8 fF μm −2), and Ca 2+ channel patches covered an average of 60% of the footprint in 11 cells. If the estimated 10 000 to 20 000 Ca 2+ channels (; ) in a chromaffin cell were distributed homogeneously throughout these patches, the average patch would contain 800–1700 channels at a density of about 25–50 μm −2. These results stand in contrast to the way Ca 2+ channels are distributed in the membrane of large synaptic terminals, such as ribbon synapses of hair cells, bipolar cells or the calyx of Held, where all channels occur in a few very tight clusters, a fraction of a micrometre in diameter, covering less than 5% of the surface membrane (;;;; ). Electron microscopy and electrophysiology studies in frog hair cells indicate that an average active zone contains about 90 Ca 2+ channels covering an area of 0.06 μm 2, equivalent to a density of 1500 channels μm −2. Calcium influx through these tight clusters of channels can be readily observed by imaging (;;; ). Excitation–secretion coupling in neuroendocrine cells and synapses The results of this study allow us to compare the membrane organization of Ca 2+ channels and fusion sites in chromaffin cells and presynaptic terminals.

Synapses provide specific sites of communication between neurons, and release of neurotransmitter occurs at active zones on the spatial scale of hundreds of nanometres directed towards postsynaptic receptors that are similarly localized. In contrast, chromaffin cells in the adrenal gland release catecholamines into the vasculature that travels along the entire apical surface of the cell, and it is across this large region of surface membrane that granules are docked. These distinctive differences in the spatial organization of exocytic sites in neurons and neuroendocrine cells might be correlated with differences in the spatial organization of Ca 2+ channels.

Larger domains of calcium entry in chromaffin cells compared to synapses may reflect the lack of spatial precision in the release of hormones compared to neurotransmitters.

Frequently Asked Questions 1. Would you please simplify the WinLTP startup? 'Putting the already defined protocols for free download would be great - e.g. One for two alternating pathway, paired-pulse field LTP recording and one for two alternating pathway, paired-pulse whole-cell LTP recording (I guess this would cover 99% of your customers).' We have refocused the default, initial startup of WinLTP back to ease of use for those researchers who just want to use WinLTP to do basic LTP/LTD experiments (hopefully less than 99%!!!). A single mouse button click of the 'MainProtocol' button starts alternating S0/S1 stimulation with signal averaging, 2.

Digidata 1322a Drivers For Mac Free

A single mouse click of the 'T0 Sweep' button evokes a single S0 train, 3. A single mouse click of the 'T1' button evokes S1 theta burst stimulation, 4. A single mouse click of the 'Repeat P0' button evokes S0 LowFrequencyStimulation to induce long-term depression. And a single mouse button click of the 'Stop' button below the 'MainProtocol' button stops the experiment, These stimulations comprise most of what an LTP/LTD experiment need Next, the alternating S0/S1 stimulation with signal averaging can be changed to just repeat S0 stimulation with signal averaging by either removing the P1sweep or unchecking the P1sweep checkbox. Or the alternating S0/S1 stimulation with signal averaging can be changed to alternating S0/S1 stimulation without signal averaging by a single mouse button click on the 'No Avg' protocol button. However, in the startup example there was only one one S0 pulse in the P0sweep, and one S1 pulse in a P1sweep.

To increase the number of pulses per sweep to 2, just change the 'Number Pulses' field from 1 to 2. And we have also included for getting started, a whole-cell patch-clamping LTP/LTD experiment with two alternating pathway S0/S1 stimulation, with signal averaging, and with paired-pulse stimulation ( PatchClampLTP.pro). 2 Will WinLTP reanalyze pop-spikes in binary multi-sweep pClamp ABF files? Yes, using Alen Eapen's, you can first convert your multi-sweep ABF files to single sweep Axon Text Files (.ATF), that WinLTP can then automatically reanalyze. Just click the 'Start Reanalysis' button and select the ATF files to reanalyze, then click OK.

WinLTP will automatically realize that the files are single sweep ATF files generated by bin2txtswps. Bin2txtswps can also convert multi-sweep binary Igor Pro IBW files and WinWCP WCP files to single sweep ATF files that WinLTP can then reanalyze. Several users have asked: What is the best way to copy over an old protocol file to a new WinLTP program when installing a new version of WinLTP? Basically what you want to do is run the old version of WinLTP with the old version protocol file alongside running the new version of WinLTP, and then manually copy the protocol field values from the old protocol file to the new one. Unfortunately, we do not have a program to automatically convert an earlier version of a protocol file to a later version of a protocol file. Here are the steps that make this conversion as straightforward as currently possible (say updating an old protocol file from WinLTP 2.01 to run in WinLTP 2.30): 1.

Copy the old WinLTP program to a new filename, e.g. Copy WinLTPm201.exe to WinLTPm201old.exe. Do NOT delete your old protocol files, i.e. Those for WinLTPm201.exe.


Uninstall the old WinLTP 2.01 program. This will include uninstalling (deleting) the old WinLTP program (i.e.

WinLTPm201.exe), but NOT the old WinLTP program that you have copied to another name (i.e. Nor will it delete the old protocol files. Install the new WinLTP 2.30 program. Start the old WinLTP program (by double-clicking WinLTPm201old.exe in the C: WinLTP folder in Windows Explorer) which will also load the last old protocol file used. Then use File-Open to load the old protocol file you wish to duplicate for the new WinLTP program. However, do NOT click the ‘MainProtocol’ button to start running this old protocol file in the old WinLTP program.

Start the new WinLTP program (by double-clicking WinLTPm230.exe in Windows Explorer) – this will NOT load the last old protocol file used, because it is the wrong size. Look at the field values in the old WinLTP program, and enter them into the new WinLTP program field values.

Save this new protocol to a new protocol file name. Then run the NEW protocol file in the NEW WinLTP program (i.e. WinLTPm230.exe) by clicking on the ‘MainProtocol’ button to see if this new protocol runs correctly. Make changes and resave as needed. It is important to run the MainProtocol of ONLY ONE of the old or new WinLTP programs AT ONE TIME.

This is because both programs are using the same data acquisition board, and running the MainProtocol in both programs would cause both programs to access the same board at the same time, and would be complete chaos. You COULD run the MainProtocol of the OLD WinLTP program when not at the same time running the MainProtocol of the NEW WinLTP program. However, it is better to only run the MainProtocol of the NEW WinLTP program because that is the new protocol file you want to test. Presumably you already know how the OLD protocol file works. How can you do voltage increment/decrement stimulations in WinLTP? WinLTP still does not have a true increment/decrement stimulation capability. However, provided the voltage pulses are not too far apart time-wise, you can put in 10 pulses of increasing amplitude in the 20 epoch stimulation for each P0/P1/T0/T1 sweep, or a total of 40 different pulse amplitudes can be delivered.

Here is a protocol which generates -80mV to +40mV in 500 msec steps every 1.5 sec. And here, PeakAmp is being measured, although other useful measurements like AvgAmp could also be measured at the end of the step (ie at steady-state). The key is to use an ‘unused’ S0 or S1 stimulation (with no connection to a Stimulus Isolator) to time when the pulse occurs so that the PeakAmp (and maybe AvgAmp) can then be analyzed as an EPSP normally would be. Is there a better way to measure electrical characteristics of neurons than WinLTP? WinLTP does a very good job of studying synaptic plasticity events like LTP, LTD etc, and is far better than programs that just tack on LTP capability as an afterthought. However, we dont pretend that WinLTP does everything. That is one of the reasons why we have always had the ' webpage on the WinLTP site.

Clearly with WinLTP lacking analog stimulation increment/decrement and P/N subtraction, there are other programs that will do a better job of analyzing electrical characteristics of cells. One solution that researchers use is to have pClamp (Molecular Devices) at the same setup as WinLTP. That works well if the researchers are using the legacy Digidata 1322A board, so that they only need one data acquisition board at their setup.

However, if pClamp users use the Digidata 1440A or 1550x boards, this requires an additional National Instruments or Digidata 1322A board for WinLTP. Thus they will need two boards at their setup. A alternative, low cost solution to better measure electrical characteristics is to use data acquisition programs that use the same National Instruments boards as WinLTP.

Some programs we would suggest trying include: 1) the free WinWCP (by John Dempster, Univ. Of Strathclyde), or 2) the low-cost AxoGraph (by John Clements, Axograph Scientific). How can I put out 900 pulses at 3 Hz stimulation? Put 3 pulses at 333.3 msec intervals into a sweep of 1 sec duration. Then output 300 of these 1 second sweeps, one every second.

7 This error is usually due to the Analog Inputs and Outputs cable being connected to a PCI or PCIexpress data acquisition board that somehow interfers with the recalibration. In the WinLTP NI board calibration dialog box we state: 'WARNING: National Instruments recommends that to accurately calibrate this board, it and the computer should warm up for at least 15 minutes, and all Analog Outputs and Inputs should be disconnected.' If you get this error, disconnect your output cable at the data acquisition board, and recalibrate. Does WinLTP run on the Apple Mac computer?

Yes for both Online/Acquisition and Reanalysis. However, neither runs as a native OSX application, but runs in Windows installed on a Mac. Apple has fairly recently converted their Mac computer and operating system to run on Intel processor / Windows compatible computers. Apple actually includes an application called BootCamp that can start (or boot) the Intel Mac computer into an user installed copy of Windows XP, Vista or 7. There is also another type of software, call virtual machine software, that can enable your Windows computer to simultaneously run Windows and Linux, or your Mac computer to run Mac OSX and Windows simultaneously along side each other. In our lab we often run the WinLTP Reanalysis program on Macs running Windows in a virtual machine created by Parallels or VMware Fusion, that runs along side OSX.

Furthermore, we have tested a National Instruments USB 2.0 M-Series board on a Mac notebook booted by BootCamp into Windows 7, and with the limited testing we have done, the board seems to run OK. We have not tested PCI or PCIexpress M- and X-Series boards or Digidata 132x boards, but it is likely that they will work too. We have not tested acquisition in a virtual machine created by Parallels or VMware Fusion and have no idea whether or not it will work.

It is highly unlikely that we will write a native WinLTP application for OSX on the Mac. Basically, we are far, far more interested in adding functionality to WinLTP rather than transferring WinLTP to another operating system. If we could just click a button on our development system to do so, we would do it, but realistically, it would probably mean a complete rewrite of WinLTP. Plus, using virtual machine operating systems like Parallels and VMware Fusion on the Mac to run Windows is the future (in fact it is here now!). Now a computer isn't a Windows or Mac or Linux computer - it's what you want, maybe all three. Will WinLTP run on any boards other than the National Instruments M- and X-Series USB/PCI/PCIexpress or the Axon Digidata 132x data acquisition boards? There are no current plans for WinLTP to run the new Axon Digidata 1440A, 1550 or 1550A data acquisition board.

And it is highly unlikely that WinLTP will run any of the CED data acquisition boards, or any of the Instrutech data acquisition boards. It's a lot of work installing a new type of data acquisition board in a multitasking program such as WinLTP. We would much rather put the work into increasing WinLTP program functionality. Also, WinLTP can never support the National Instruments E-Series data acquisition boards because they do not have streaming digital output.

To us the M- and X-series boards for the USB 2.0, PCI and PCIexpress busses are functionally extremely good with WinLTP and come at a very reasonable price. In addition, the M- and X-Series boards can run some other very good data acquisition programs including:, and programs written in,. What are the advantages of the M- and X-Series boards versus the Digidata 132x boards? A) You can still buy the M- and X-Series boards from National Instruments and will be able to for a long time into the future. Molecular Devices no longer sells the legacy Digidata 132x boards. B) The delay in response to keyboard input for altering protocol values is much less for the M- and X-Series PCI/PCIe boards ( 0.5 sec) compared to the Digidata 132x boards ( 5.0 sec).

Will WinLTP's programmer be willing to write any scripts in the Protocol Builder? Yes, within reason. We know it can be extremely tough when trying to write scripts for a system you don't know anything about, so I'm glad to get users started in script programming. Plus, the script programming component in the Protocol Builder is the part of WinLTP 'm most proud about (along with the multitasking - the rest is pretty much grunt work). And we think that the WinLTP scripting is bettered by few electrophysiology data acquisition systems.