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Abstract

  1. Top of page
  2. Abstract
  3. METHODS
  4. RESULTS
  5. DISCUSSION
  6. Acknowledgements
  • We used the patch-clamp technique, in conjunction with membrane capacitance measurement, fluorescence measurement of intracellular calcium concentration ([Ca2+]i), and flash photolysis of caged Ca2+ to study exo- and endocytosis in identified rat corticotrophs.

  • Exocytosis stimulated by depolarization pulses was typically followed by a ‘slow’ endocytosis that retrieved the membrane with a time constant of ∼6 s. The efficiency (the endocytosis/exocytosis amplitude ratio) of ‘slow’ endocytosis was ∼1.2 at [Ca2+]i < 3 μm and increased to ∼1.6 at [Ca2+]i > 3 μm.

  • Whole-cell dialysis through a patch pipette did not affect the kinetics and the efficiency of ‘slow’ endocytosis, but the amplitude of exocytosis was reduced.

  • ‘Slow’ endocytosis did not require sustained [Ca2+]i elevation and its kinetics was only weakly [Ca2+]i dependent. Our results suggest that ‘slow’ endocytosis involves a Ca2+ sensor with a high Ca2+ affinity (∼500 nm).

  • At high [Ca2+]i (> 10 μm), the ‘slow’ endocytosis was frequently preceded by a ‘fast’ endocytosis that comprised multiple steps of rapid decrease in membrane capacitance.

  • Neither calmodulin nor calcineurin appeared to be the Ca2+ sensor for endocytosis because the two forms of endocytosis were not affected by the calmodulin inhibitor calmidazolium (500 μm) or the calcineurin inhibitors cyclosporin A (1 μm) and calcineurin autoinhibitory peptide (1 mg ml−1). Ba2+, a poor activator of calmodulin, could support both forms of endocytosis but slowed the kinetics of ‘slow’ endocytosis ∼2-fold.

  • Non-hydrolysable analogues of GTP (GDP-β-S) and ATP (ATP-γ-S) also failed to inhibit either form of endocytosis, indicating that neither GTP nor ATP was essential for endocytosis.

  • We suggest that the high Ca2+ affinity of ‘slow’ endocytosis may be important for maintaining continuous cycles of exocytosis-endocytosis during sustained adrenocorticotropin secretion in corticotrophs.

The major function of corticotrophs is the secretion of the stress hormone, adrenocorticotropin (ACTH). Similar to the secretion of other hormones and neurotransmitters, the secretion of ACTH is Ca2+ dependent and involves the process of exocytosis. During exocytosis, the secretory vesicle membrane is fused with the cell plasma membrane and the contents of the vesicles are released. As a corollary, the reverse of exocytosis, endocytosis must also occur, because the cell plasma membrane does not increase monotonically. The coupling between exocytosis and endocytosis is demonstrated most clearly in synapses where fluorescent dyes have been used to label vesicle membrane through multiple rounds of exocytosis and endocytosis (Betz & Bewick, 1992; Ryan et al. 1996; Wu & Betz, 1996; Klingauf et al. 1998). In non-neuronal cells, such as melanotrophs and chromaffin cells, endocytosis following stimulated exocytosis has been studied using the patch-clamp method and membrane capacitance measurement (Thomas et al. 1994; Artalejo et al. 1996; Smith & Neher, 1997; Engisch & Nowycky, 1998; Nucifora & Fox, 1998). These studies show that endocytosis can occur shortly after stimulated exocytosis, but it is often variable. Endocytosis also appears to be sensitive to dialysis or loss of small soluble molecules through the patch pipette in the whole-cell configuration (Eliasson et al. 1996; Smith & Neher, 1997). In chromaffin cells and melanotrophs, the time constant of endocytosis for retrieving membrane after stimulated exocytosis ranged from 5 to 15 s. A faster form of endocytosis with a time constant of less than a second has also been described in these cells (Neher & Zucker, 1993; Thomas et al. 1994), as well as in the nerve terminals of the posterior pituitary (Hsu & Jackson, 1996). This fast endocytosis has been attributed to the retrieval of membrane added by earlier rounds of exocytosis (Thomas et al. 1994; Smith & Neher, 1997; but see Engisch & Nowycky, 1998). In melanotrophs and chromaffin cells, endocytosis has been suggested to be Ca2+ dependent (Thomas et al. 1994; Smith & Neher, 1997; Engisch & Nowycky, 1998). However, in nerve terminals, endocytosis has been suggested to be Ca2+ independent (Wu & Betz, 1996), and high [Ca2+]i has been reported to inhibit endocytosis (von Gersdorff & Matthews, 1994; Hsu & Jackson, 1996). Here, we take advantage of the robust endocytosis following stimulated exocytosis in rat corticotrophs to study the relationship between [Ca2+]i, endocytosis and exocytosis. Using the patch-clamp technique along with membrane capacitance measurement, [Ca2+]i measurement with indo-1 fluorescence, and flash photolysis of caged Ca2+, we find that endocytosis in rat corticotrophs involves a Ca2+ sensor with high Ca2+ affinity (≈500 nm), and that [Ca2+]i may directly modulate the amount of endocytosis.

METHODS

  1. Top of page
  2. Abstract
  3. METHODS
  4. RESULTS
  5. DISCUSSION
  6. Acknowledgements

Cell preparation

The anterior lobe of the pituitary glands were removed from a male Sprague-Dawley rat (age 5-6 weeks) killed with an overdose of halothane in accordance with the standards of the Canadian Council on Animal Care. Anterior pituitary glands were dissociated enzymatically using collagenase and trypsin as previously described (Tse & Hille, 1994). Single corticotrophs were identified from the heterogeneous population by reverse haemolytic plaque assay (Smith et al. 1984). The procedures were similar to that described previously (Lee & Tse, 1997; Tse & Tse, 1998). Briefly, the dissociated pituitary cells were suspended in Dulbecco's modified Eagle's medium (DMEM; Gibco, Grand Island, NY, USA) that contained 0.1 % (w/v) bovine serum albumin (BSA, Sigma, St Louis, MO, USA). The pituitary cell suspension was then mixed with one-third the volume of 12 % (v/v) sheep erythrocytes (Colorado Serum Co., Denver, CO, USA) in 0.9 % (w/v) NaCl. The erythrocytes were previously conjugated with Staphylococcus aureus-derived protein A (Sigma), using 0.2 mg ml−1 CrCl3 as a catalyst. The cell mixture was incubated with a mixture of 10 nm corticotropin-releasing hormone (CRH) and 100 nm arginine vasopressin (AVP), and rabbit polyclonal antibodies to rat ACTH (1:20 dilution; gift from Dr R. J. Kemppainen, Auburn University, Auburn, AL, USA) for 3 h at 37 °C. Plaques were formed by a 30 min exposure to guinea-pig complement at 1:50 dilution. The cells were maintained under standard culture condition in a DMEM medium supplemented with 10 % (v/v) horse serum, 50 U ml−1 penicillin G and 50 μg ml−1 streptomycin. Recordings were made in cells cultured for 2-4 days after plaque formation.

Solutions

The standard bath solution contained (mm): 150 NaCl, 10 Hepes, 8 glucose, 2.5 KCl, 5 or 10 CaCl2 and 1 MgCl2 at pH 7.4 (titrated with NaOH). Apamin (0.5 μm) was also added to the bath solution to block the SK-type Ca2+-activated K+ channels. In experiments where Ba2+ was used to trigger exocytosis, Ca2+ in the standard bath solution was replaced with 5 mm Ba2+. The standard pipette solution contained (mm): 70 caesium aspartate, 20 tetraethylammonium chloride, 40 Hepes, 1 MgCl2, 2 Na2ATP and 0.1 Na4GTP at pH 7.4 (titrated with CsOH). In experiments where [Ca2+]i was elevated via flash photolysis of caged Ca2+, 10 mm nitrophenyl-EGTA (NP-EGTA; 80-95 % saturated with Ca2+) was included in the pipette solution.

Stocks of AVP and CRH (Peninsula Laboratories, Belmont, CA, USA) were dissolved in 0.1 m acetic acid, lyophilized and kept at -20 °C. Indo-1 (Calbiochem, La Jolla, CA, USA), indo-1 FF (Texas Fluorescence Labs, Inc., Austin, TX, USA) and NP-EGTA (gift from Dr G. Ellis-Davis, MCP Hahnemann University, PA, USA) were kept as stock solutions in distilled water at -20 °C. Calmidazolium and cyclosporin A (Calbiochem) were dissolved in DMSO and kept as stock solution at -20 °C. GTP-γ-S, GDP-β-S, ATP-γ-S and calcineurin autoinhibitory peptide were also obtained from Calbiochem and apamin was obtained from Sigma.

Electrophysiology

Single corticotrophs identified with the reverse haemolytic plaque assay were surrounded by a zone of lysed erythrocytes (plaque) and had no electrical coupling with any other cells. Corticotrophs are small (10-15 μm in diameter), round cells with a membrane capacitance of 3-5 pF and an input resistance of ≈5 GΩ (Lee & Tse, 1997). In this study, individual corticotrophs were voltage clamped with the whole-cell gigaseal method (Hamill et al. 1981) using an EPC-7 patch-clamp amplifier. After the establishment of the whole-cell configuration, the series resistance of the pipette and the membrane capacitance of the cell were neutralized by the patch-clamp amplifier and the cell was voltage clamped at a holding potential of -90 mV. Membrane capacitance measurement was made by superimposing an 800 Hz, 30 mV peak-to-peak sinusoidal voltage onto the holding potential, and then separating the resultant current with a phase lock-in amplifier as described previously (Tse & Hille, 1994). To minimize conductance changes during capacitance measurements, the K+ channel blockers Cs+ and TEA+ were included in the pipette solution and apamin was included in the bath to inhibit the SK-type Ca2+-activated K+ channels in these cells (Tse & Lee, 1998). Under this experimental condition, changes in capacitance were not accompanied by any corresponding changes in conductance. Throughout the time period of capacitance change (< 30 s), there is minimal change in conductance (< 0.5 nS). Capacitance and conductance traces were first recorded on VCR tapes using a NeuroData PCM recorder (Neuro Data Instruments Corp., New York), and digitized later. Holding potential and voltage pulses were controlled using an IBM-compatible PC and the data acquisition program pCLAMP v.6 (Axon Instruments, Foster City, CA, USA). The pipettes were made from haematocrit glass (VWR Scientific Canada Ltd, London, Ontario, Canada) and the resistance was 2-4 MΩ after filling and 5-15 MΩ during whole-cell recording. All experiments were performed at room temperature (20-23 °C). A -10 mV junction potential was corrected throughout. Values given in the text are means ±s.e.m. To photolyse Ca2+-NP-EGTA, a UV flash from a modified XF-10 xenon flash lamp (Hi-Tech Ltd, Salisbury, UK) was delivered to the cell via a fused silica focusing lens which replaced the microscope's condenser (Tse et al. 1997). Different [Ca2+]i levels were achieved by varying the percentage of Ca2+ load in the NP-EGTA solution or the intensity of the UV flash.

[Ca2+]i measurement

In experiments where exocytosis was stimulated with depolarizations, [Ca2+]i measurement was made with the Ca2+-sensitive dye indo-1. In experiments where exocytosis was stimulated with flash photolysis of caged Ca2+, indo-1 FF, a Ca2+-sensitive dye with a lower Ca2+ affinity was used. Indo-1 or indo-1 FF (0.1 mm) was included in the pipette solution and loaded into the cell via the whole-cell patch pipette. Details of the instrumentation and procedures of [Ca2+]i measurement were as described previously (Tse et al. 1994; Lee & Tse, 1997; Tse & Tse, 1998). Briefly, the dye was excited by 365 nm light delivered from a HBO 100 W mercury lamp via a × 40, 1.3 NA UV fluor oil objective lens (Nikon). Emission fluorescence at 405 and 500 nm was collected with two photomultiplier tubes (Hamamatsu H3460-04). The output of the photomultiplier tubes was converted to TTL pulses, and counted by a CYCTM-10 counter card (Cyber Research Inc., Branford, CT, USA) installed in an IBM-compatible PC. [Ca2+]i was calculated from the ratio of the two emission fluorescence measurements (R) according to the equation of Grynkiewicz et al. (1985):

  • image(1)

where Rmin is the ratio when Ca2+ is strongly chelated with EGTA, Rmax is the ratio when 15 mm free Ca2+ is present, and K* is a constant determined empirically. Rmin was measured in cells loaded with (mm): 52 potassium asparate, 50 Hepes, 50 EGTA, and 10 KCl (titrated to pH 7.4 with KOH). Rmax was measured in cells loaded with (mm): 136 potassium asparate, 50 Hepes, and 15 CaCl2 (titrated to pH 7.4 with KOH). For indo-1 measurement, K* was calculated from eqn (1) using R values obtained from cells loaded with (mm): 60 potassium aspartate, 50 K-Hepes, 20 K-EGTA, 15 CaCl2, 0.1 indo-1 (pH 7.4), which had a calculated free Ca2+ concentration of 212 nm at 24 °C (Blinks et al. 1982). For indo-1 FF measurements, K* was estimated from cells loaded with (mm): 15 potassium aspartate, 70 diglycolic acid, 50 Hepes and 10 CaCl2 (titrated to pH 7.4 with KOH), which had a free Ca2+ concentration of 24 μm (measured with a Ca2+ electrode). In the presence of Ca2+-NP-EGTA compounds, the value of Rmin was obtained individually for each cell as the average of 5-10 measurements taken immediately before the flash.

Data analysis

The change in membrane capacitance was analysed using Origin 4.1 (Microcal Software, Northampton, MA, USA). The capacitance trace was adjusted for baseline drift by subtracting a straight line that was fitted to at least 2 s of the capacitance trace preceding the stimulus (depolarization pulse train or flash photolysis of caged Ca2+). On average, the drift in the baseline of the capacitance trace was 0.33 ± 0.03 fF s−1 (n = 100). The stimulus triggered an increase in membrane capacitance (reflecting exocytosis) that was followed by a decay (reflecting endocytosis). In most experiments with a depolarization pulse train as stimulus, only ‘slow’ endocytosis was observed. For ‘slow’ endocytosis, the decay of the capacitance trace could be described by a single exponential. The fit of the exponential generally started from the steepest part of the capacitance decay and ended at the baseline. The amplitude of endocytosis was taken from the net decrease in capacitance that was fitted by the exponential. In cells that exhibited both ‘fast’ and ‘slow’ endocytosis, the exponential fit for ‘slow’ endocytosis started from where the ‘fast’ endocytosis apparently ended. For exocytosis stimulated with flash photolysis of caged Ca2+, the UV flash created a transient artifact that contaminated the capacitance trace immediately after the flash. The rising phase of this artifact takes about 2 ms and the falling phase could be described as the sum of two decaying exponentials with time constants of 0.16 s and 5 s. The amplitude of the slower exponential is described by Aslow= 2 + 0.17Atotal, where Atotal is the peak amplitude of the artifact or the sum of the initial amplitudes of the two exponentials. To correct for this artifact, the two decaying exponentials (scaled for the peak of the artifact) were subtracted from the membrane capacitance trace. All values of rate of exocytosis and amount of exocytosis or endocytosis were normalized to the initial cell size and then multiplied by the average cell size of 3.4 pF.

Kinetic model

Assuming that exocytosis and endocytosis are consecutively coupled, they can be described by the following model:

  • image

where A is the readily releasable vesicles, B is the exocytosed vesicles and C is the endocytosed vesicles. k1 is the rate constant of exocytosis and k2 is the rate constant of endocytosis. The kinetics of these three types of vesicles can be represented by the following equations:

  • image(2)
  • image(3)
  • image(4)

where CA(t), CB(t) and CC(t) are the pool sizes of A, B and C at time t. CA(0) is the pool size of A at time zero or the pool size of the readily releasable vesicles. To generate the simulations (continuous lines) in Fig. 6a and B, CA(0) was set to 90 fF (estimated from the maximal exocytic response in Fig. 6a), k2 was set to 0.35 s−1 (τ of 2.86 s; estimated from Fig. 5B) and k1 was set to 1.5 × the rate of exocytosis/CA(0) (the scaling factor of 1.5 was applied to adjust for the lower estimate of the rate of exocytosis). Since the average efficiency of endocytosis (endocytosis/exocytosis amplitude) was ≈1.2, CC(t) was scaled up by 1.2 to allow for excess endocytosis. Then the new equation for CB(t) is:

  • image(5)

CB(t) reaches its maximum or peak when dCB(t)/dt= 0, and solving for t we get (ln[{k1+ 0.2k2}/1.2k2])/(k1 - k2) which is the time of the peak. The maximum size of CB(t) is then calculated by using this value of t and eqn (5).

image

Figure 6. Figure 6. Cumulative exocytosis and the time to the peak of exocytosis are dependent on the rate of exocytosis

A, plot of cumulative exocytosis versus the rate of exocytosis. Each data point is from individual cells that were stimulated by flash photolysis of caged Ca2+. ▪, cells where only ‘slow’ endocytosis was present; □, cells where both ‘fast’ and ‘slow’ endocytosis were present. Cumulative exocytosis increased with the rate of exocytosis and approached ∼90 fF at high rates of exocytosis. B, plot of the time to the peak of exocytosis versus the rate of exocytosis. The time to the peak of exocytosis is the time interval between the UV flash and the peak of the Cm trace. Same data set as in A is used. In both A and B, the continuous lines are obtained from simulations of a kinetic model which assumes that endocytosis follows immediately the exocytosis of individual granules (see Methods).

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image

Figure 5. Kinetics of ‘slow’ endocytosis at high [Ca2+]i

A, recordings from the same three cells shown in Fig. 4A are shown on a 10-fold slower time scale. Exocytosis was followed by endocytosis. [Ca2+]i decreased slowly after the rapid rise stimulated by flash photolysis of caged Ca2+. τendo was derived from the fit of a single decaying exponential to the Cm decrease. Note that for the record in the rightmost panel, ‘slow’ endocytosis was preceded by a ‘fast’ endocytosis. The τendo for this cell was derived from the fit of a single decaying exponential to the ‘slow’ component only of the Cm decrease. B, plot of τendoversus peak [Ca2+]i. Data for individual cells (□) and their grouped averages (▪) are shown. The rate constant is the inverse of τendo.

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Numerical simulation of endocytosis

We numerically simulated endocytosis during the [Ca2+]i recovery after a depolarization pulse train had raised [Ca2+]i. The recovery of [Ca2+]i is described by a single exponential with a time constant of 2.3 s. The rate constant of endocytosis, k, is Ca2+ dependent and is described by the logistic equation:

  • image(6)

where the maximum rate constant is set to 0.35 (estimated from the rate constant at 15 μm[Ca2+]i in Fig. 5B), and the half-maximal value of k is set to 500 nm[Ca2+]i. To describe endocytosis, we use an iterative expression:

  • image(7)

where Ai is the amplitude of endocytosis and ki is the rate constant of endocytosis at time step i. The time step interval is 0.2 s. Since k is a function of [Ca2+]i, ki is calculated from the corresponding [Ca2+]i at time step i with eqn (6). To obtain the time constant of ‘slow’ endocytosis (τendo) from the simulated endocytosis trace, we fitted a single exponential to the simulated endocytosis trace.

RESULTS

  1. Top of page
  2. Abstract
  3. METHODS
  4. RESULTS
  5. DISCUSSION
  6. Acknowledgements

Endocytosis following depolarization-triggered exocytosis

In corticotrophs, exocytosis was typically followed by membrane retrieval (endocytosis). To determine if endocytosis was affected by whole-cell dialysis, we stimulated single corticotrophs at different times after the establishment of whole-cell recording. One such experiment is shown in Fig. 1a. The cell was held at -90 mV and [Ca2+]i was monitored via indo-1. At 5, 10.5 and 17 min after whole-cell recording, a train of five depolarizing voltage steps (each 500 ms in duration) to +10 mV was delivered at 1.25 Hz to activate voltage-gated Ca2+ channels. Each depolarization train stimulated a [Ca2+]i rise that was accompanied by a membrane capacitance (Cm) increase or exocytosis. Following the stimulus, [Ca2+]i quickly recovered to pre-stimulus levels, while Cm decayed gradually, reflecting endocytosis. In this cell, although the three trains of depolarization triggered a similar rise in [Ca2+]i, the exocytic response diminished with the time of whole-cell dialysis. Unlike exocytosis, endocytosis persisted throughout the time course of the experiment and was not affected by whole-cell dialysis. Similar experiments were repeated in seven cells and the amount of exocytosis (normalized to the exocytosis triggered at 5 min after whole-cell in individual cells) was plotted against the time after the establishment of whole-cell recording in Fig. 1B. In these cells, the exocytic response diminished with the duration of whole-cell dialysis, while the depolarization-triggered [Ca2+]i elevation was mostly unchanged (< 5 %). This run-down in exocytic response might be attributed to a gradual loss of small soluble molecules (via the whole-cell pipette) that were important for the replenishment of the readily releasable pool of vesicles. Figure 1B also shows a plot of the efficiency of endocytosis (the endocytosis/exocytosis amplitude ratio) versus the duration of whole-cell dialysis. Unlike exocytosis, the efficiency of endocytosis was not affected by whole-cell dialysis. In the example shown in Fig. 1a, as well as in most experiments where exocytosis was stimulated with depolarization pulses, the time course of endocytosis (decay of Cm) could be described by a single decaying exponential. In the following sections, we shall refer to this form of endocytosis as ‘slow’ endocytosis. Figure 1C shows that whole-cell dialysis also had little effect on the time constant of ‘slow’ endocytosis (τendo). These results suggest that ‘slow’ endocytosis is strongly coupled to exocytosis, but unlike exocytosis, the maintenance of ‘slow’ endocytosis is not affected by whole-cell dialysis.

image

Figure 1. Endocytosis does not wash out with whole-cell dialysis

A, a corticotroph was voltage clamped at -90 mV and stimulated with five 500 ms pulses to 10 mV at 1.25 Hz at the indicated time after establishment of whole-cell recording. Exocytosis and endocytosis were measured as the increase and decrease respectively in the membrane capacitance (Cm). Simultaneous measurement of [Ca2+]i is also shown. B, exocytosis, but not the efficiency of ‘slow’ endocytosis, runs down with whole-cell dialysis. The amplitude of exocytosis (ΔCm) is normalized to the value at 5 min and is plotted against the time after establishment of whole-cell recording (▪). The efficiency of endocytosis (‘slow’ endocytosis/exocytosis amplitude ratio) is shown in the same plot (□). C, the kinetic of ‘slow’ endocytosis is not affected by whole-cell dialysis. The time constant of endocytosis (τendo) derived from the fit of a single decaying exponential to the Cm decrease is plotted against the time after establishment of whole-cell recording. The dotted line indicates the average τendo (6.7 ± 0.3 s) of all the data points. The numbers in parentheses indicate the number of cells averaged for each data point. The same data set was used in B and C.

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Figure 1B shows that the average efficiency of ‘slow’ endocytosis had a value greater than one. This implied that more membrane was retrieved than was added by exocytosis, or excess membrane retrieval. Figure 2a shows records from three cells that exhibited different amounts of excess membrane retrieval. Higher [Ca2+]i appeared to trigger more excess membrane retrieval. In Fig. 2B, we plotted the efficiency of ‘slow’ endocytosis against the amplitude of the Ca2+ trigger. At [Ca2+]i < 3.5 μm, the efficiency of ‘slow’ endocytosis was ≈1.2 and it increased to ≈1.7 at 5 μm[Ca2+]i. To examine whether more exocytosis also stimulated more endocytosis, we plotted the efficiency of endocytosis against the amplitude of exocytosis in Fig. 2C. The efficiency of ‘slow’ endocytosis was not strongly influenced by the amount of exocytosis. These results suggest that [Ca2+]i may directly modulate the efficiency of endocytosis, and thus the amount of endocytosis.

image

Figure 2. High [Ca2+]i stimulates more excess membrane retrieval

A, examples of three different cells stimulated by voltage pulses to different [Ca2+]i levels are shown. Note that the amplitude of exocytosis is similar for the three cells, but the efficiency of ‘slow’ endocytosis is different. B, the efficiency of ‘slow’ endocytosis is plotted against the peak [Ca2+]i that stimulated each exo/endocytotic event. The efficiency was higher at higher [Ca2+]i. C, plot of the efficiency of endocytosis (Endo: exo ratio) versus the amplitude of exocytosis. The efficiency of ‘slow’ endocytosis was not influenced by the amount of exocytosis. The numbers in parentheses indicate the number of exo/endocytotic events averaged for each data point.

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Ca2+ dependence of endocytosis

Since endocytosis was coupled to exocytosis and exocytosis was Ca2+ dependent, the Ca2+ that stimulated exocytosis also might regulate the kinetics of endocytosis. Note that in Fig. 2a, however, endocytosis clearly continued after [Ca2+]i had returned to pre-stimulus levels. Following the depolarization stimulus, [Ca2+]i decayed with an average time constant of 2.3 ± 0.1 s (n = 79), almost 3 times faster than the time constant of ‘slow’ endocytosis (6.1 ± 0.2 s, n = 96). This suggests that after endocytosis is triggered, the completion of endocytosis does not require an elevated [Ca2+]i. Nevertheless, the amplitude of the Ca2+ trigger may still affect the kinetics of endocytosis. To address this, we examined exo- and endocytotic events triggered by different levels of [Ca2+]i elevation. In the example shown in Fig. 3a, two different depolarization stimuli were delivered to the same cell at ≈5 min apart. The first stimulus, a train of three 500 ms pulses at 1.25 Hz, elicited a larger [Ca2+]i elevation and triggered more exocytosis than the second stimulus, which comprised a single 500 ms pulse. Note that the endocytosis following the first stimulus consisted of an initial rapid decrease that comprised multiple steps (see for example, Fig. 7B and D for an expanded time scale) and was followed by the more typical ‘slow’ endocytosis. We shall refer to these rapid step-like decreases in capacitance as ‘fast’ endocytosis, which will be described later. For ‘slow’ endocytosis, τendo was similar for both small and large [Ca2+]i elevations (4.3 s vs. 4.0 s). In another set of experiments, the amount of indo-1 was increased from 100 to 500 μm (Fig. 3B). The extra indo-1 reduced the peak of the [Ca2+]i elevation and slowed the [Ca2+]i recovery after the stimulus. Nevertheless, the τendo for the two very different [Ca2+]i stimuli in Fig. 3B was similar (6.2 s vs. 5.4 s). Figure 3C summarizes the results from the the two sets of experiments in Fig. 3a and B. The values of τendo decreased slightly with higher [Ca2+]i elevations for both experiments with 100 μm and 500 μm indo-1. At ≈5 μm[Ca2+]i, τendo was 5.2 ± 0.3 s (n = 18), while at ≈2 μm[Ca2+]i, τendo was 6.4 ± 0.5 s (n = 20). Thus, the rate of endocytosis is only weakly influenced by the amplitude of the Ca2+ trigger.

image

Figure 3. Influence of [Ca2+]i on the kinetics of ‘slow’ endocytosis

A shows two exo/endocytotic events from a cell recorded with 100 μm indo-1. The first event was stimulated with three 500 ms pulses at 1.25 Hz, while the second event was stimulated with a single 500 ms pulse. ‘Slow’ endocytosis during the first event was preceded by rapid decreases in Cm or ‘fast’ endocytosis. These rapid decreases were excluded from the exponential fit that gives τendo. Although the amplitude of the [Ca2+]i elevation as well as the amount of exocytosis was smaller during the second stimulation, the time constants of ‘slow’ endocytosis (τendo) in both events were similar. B shows two exo/endocytotic events from two different cells recorded with 500 μm indo-1. The τendo values were similar despite the difference in [Ca2+]i elevation. Note the slower kinetics of [Ca2+]i recovery with 500 μm indo-1. C, plot of τendoversus peak [Ca2+]i showing that τendo was not strongly influenced by [Ca2+]i. The numbers in parentheses indicate the number of exo/endocytotic events recorded with 100 μm indo-1 for each data point, and the numbers in brackets indicate the number of exo/endocytotic events recorded with 500 μm indo-1 for each data point.

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image

Figure 7. ‘Fast’ endocytosis contains step-like decreases in Cm

A, ‘fast’ and ‘slow’ endocytosis in a cell stimulated with depolarization pulses. The initial part of the Cm decrease is dominated by ‘fast’ endocytosis while the later part is ‘slow’ endocytosis. B, ‘fast’ endocytosis and the corresponding [Ca2+]i from A are shown on an expanded time scale. Note the large and rapid downward step-like decrease in Cm (Cm scale has also been expanded). C, ‘fast’ and ‘slow’ endocytosis in a cell stimulated with flash photolysis of caged Ca2+. D, ‘fast’ endocytosis and the corresponding [Ca2+]i from C are shown on an expanded time scale. The arrow indicates the time at which the UV light flash was applied.

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Relationship between exocytosis and endocytosis

In order to examine more closely the relationship between exocytosis and endocytosis, we used flash photolysis of caged Ca2+ to raise [Ca2+]i uniformly in the cell, thus avoiding the problem of the [Ca2+]i gradient which was inherent in the depolarization stimulus protocol. By varying the intensity of the UV flash or the Ca2+ loading of NP-EGTA, [Ca2+]i could be raised to different levels. Figure 4a shows exocytosis from three different cells when [Ca2+]i was raised to different levels via flash photolysis of caged Ca2+. A flash of UV light was delivered at the time indicated by the arrow. The flash rapidly raised [Ca2+]i which in turn stimulated exocytosis. Higher [Ca2+]i stimulated faster exocytosis. Figure 4B shows a plot of the rate of exocytosis (ΔCmt, estimated from the slope of a line fitted to the Cm trace when Cm increased from 5 to 20 fF) at different [Ca2+]i levels. The plot shows the rate of exocytosis from individual cells, as well as their grouped averages. For [Ca2+]i between 8 and 15 μm, the average rate of exocytosis increased nearly 6-fold. Thus, the rate of exocytosis is strongly dependent on the amplitude of [Ca2+]i elevation.

image

Figure 4. Exocytosis is steeply dependent on [Ca2+]i

A, flash photolysis of caged Ca2+ raised [Ca2+]i and stimulated exocytosis in three different cells. [Ca2+]i was measured with indo-1 FF. The rate of exocytosis was estimated from the slope of a line fitted to the Cm trace when Cm increased from 5 to 20 fF. The arrow indicates the time at which the flash of UV light was applied. B, plot of the rate of exocytosis versus peak [Ca2+]i using data derived from experiments similar to those shown in A. Data from individual cells (□) and their grouped averages (▪) are shown. Note that, at the range of [Ca2+]i shown here, the rate of exocytosis increased ≈6-fold. The rate constant is obtained by dividing the rate of exocytosis by 90 fF, an estimate of the size of the readily releasable pool of vesicles.

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For each cell shown in Fig. 4a, exocytosis was followed by endocytosis. Figure 5a shows the same three cells on a 10-fold slower time scale. In the records shown in the left and middle panels of Fig. 5a, endocytosis could be described by the fit of a single decaying exponential to the Cm decrease like the ‘slow’ endocytosis described above. The Cm record shown in the rightmost panel of Fig. 5a was similar to that shown in Fig. 3a; endocytosis comprised a rapid decrease in Cm that was followed by a slow component. The slow component reflected ‘slow’ endocytosis and could be described by a single exponential fit. The rapid decrease in Cm reflected ‘fast’ endocytosis which will be discussed later (see below), and was excluded from the exponential fit that gave τendo. In Fig. 5B, we plotted the τendo obtained at different [Ca2+]i from individual cells, as well as their grouped averages (the same group of cells shown in Fig. 4B). The [Ca2+]i elevation in these caged Ca2+ experiments was higher than that recorded during the depolarization experiments (Fig. 3C). Consistent with the observations illustrated in Fig. 3C, Fig. 5B shows that the rate of endocytosis increased slightly with [Ca2+]i. At 9 μm[Ca2+]i, τendo was 3.9 ± 0.8 s (n = 6) (cf. 5.2 ± 0.3 s at 5 μm in Fig. 3C), and at 15 μm, τendo decreased by ≈40 % to 2.4 ± 0.7 s (n = 4). A comparison of Fig. 4B and Fig. 5B shows that exocytosis was a faster process than endocytosis. At [Ca2+]i of ≈9 μm, the average rate constant of exocytosis (0.77 ± 0.31 s−1; Fig. 4B) was about 3-fold larger than that of endocytosis (0.26 ± 0.06 s−1; Fig. 5B). At a [Ca2+]i of ≈15 μm, the average rate constant of exocytosis (5.0 ± 0.6 s−1; Fig. 4B) was about 10-fold greater than the rate constant of endocytosis (0.42 ± 0.12 s−1; Fig. 5B). Exocytosis is evidently more steeply [Ca2+]i dependent than endocytosis.

We noted that when corticotrophs were stimulated by low [Ca2+]i elevations (< 12 μm) (e.g. left panel of Fig. 4a), cumulative exocytosis was typically small and exocytosis apparently stopped while [Ca2+]i remained elevated. To examine this further, we plotted cumulative exocytosis against the rate of exocytosis for individual cells from experiments similar to those shown in Fig. 4a. Figure 6a shows that cumulative exocytosis increased with the rate of exocytosis. When the rate of exocytosis was high, cumulative exocytosis approached 90 fF, which was approximately the size of the readily releasable pool of vesicles in corticotrophs (Tse & Tse, 1998). Thus, when the rate of exocytosis was high, cumulative exocytosis was limited by the size of the readily releasable pool of granules. However, at lower rates of exocytosis, cumulative exocytosis was less than the size of the readily releasable pool of granules. Since [Ca2+]i was elevated for many seconds after the UV flash, exocytosis should continue to occur until the readily releasable pool is exhausted. One possible explanation for this is that endocytosis had started to occur while exocytosis was still continuing and the cumulative exocytosis in our measurement was the net result of the two processes. At low [Ca2+]i, the rate constants of exocytosis and endocytosis were similar (Fig. 4B and Fig. 5B), so their mutual interference was probably greatest. At high [Ca2+]i, however, the rate constant of exocytosis was many times larger than the rate constant of endocytosis, and exocytosis would be almost complete before much endocytosis occurred, so there would be little mutual interference. To test this hypothesis, we simulated a kinetic model:

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where A represented the readily releasable vesicles, B the exocytosed vesicles and C the endocytosed vesicles. We estimated the rate constant of exocytosis (k1) from the measured rate of exocytosis, and assumed a fixed rate constant of endocytosis (k2) (see Methods). We found that this kinetic model predicted a relationship between cumulative exocytosis and the rate of exocytosis (continuous line) that was compatible with the experimental result (symbols) shown in Fig. 6a. The same kinetic model also predicted that the time for exocytosis to reach its peak - the time between the UV flash and the peak of the Cm trace - should shorten with increasing rate of exocytosis. Figure 6B shows that a shorter time to peak was associated with a faster rate of exocytosis, and the predicted relationship (continuous line) from the kinetic model was also compatible with this experimental result (symbols).

Note that in most of the experiments which exhibited a high rate of exocytosis ‘fast’ endocytosis was also present in addition to ‘slow’ endocytosis (□ in Fig. 6a and B), but it did not affect the relationships between cumulative exocytosis and rate of exocytosis, or the time to peak and rate of exocytosis. ‘Fast’ endocytosis therefore did not appear to interfere with the close coupling between ‘slow’ endocytosis and exocytosis. This observation also suggested that the initiation of ‘fast’ endocytosis was slow relative to exocytosis, so that cumulative exocytosis was not diminished by ‘fast’ endocytosis any more than by ‘slow’ endocytosis alone. It raises the possibility that ‘fast’ endocytosis may also be coupled to exocytosis in some way.

Fast endocytosis

In some corticotrophs, ‘slow’ endocytosis was preceded by a ‘fast’ endocytosis (e.g. Fig. 3a and Fig. 5a). While ‘slow’ endocytosis had a τendo of several seconds, ‘fast’ endocytosis comprised multiple step-like decreases in capacitance. ‘Fast’ endocytosis occurred occasionally in cells stimulated by depolarization pulses, and more frequently in cells stimulated with flash photolysis of caged Ca2+. Figure 7 shows an example of ‘fast’ endocytosis in a cell stimulated with depolarization pulses (Fig. 7a) and in a cell stimulated with flash photolysis of caged Ca2+ (Fig. 7C). Note that the decrease in Cm during ‘fast’ endocytosis was not smooth, but was characterized by one or more large rapid downward steps. The size of the downward steps ranged from 5 to 75 fF, corresponding to vesicles with diameters ranging from 0.4 to 1.5 μm (assuming a specific membrane capacitance of 10 fF μm−2). In all cells that exhibited ‘fast’ endocytosis, the ‘fast’ endocytosis was always followed by ‘slow’ endocytosis. ‘Fast’ endocytosis has been postulated to retrieve membrane from earlier rounds of exocytosis (Thomas et al. 1994; Smith & Neher, 1997; but see Engisch & Nowycky, 1998). Therefore, we examined whether the cell size of corticotrophs showing ‘fast’ endocytosis was bigger than that of cells that did not show ‘fast’ endocytosis. From our depolarization experiments, we found that for cells with ‘fast’ endocytosis, the cell size ranged from 2.8 to 4.2 pF (mean = 3.4 ± 0.1 pF; n = 13). For cells without ‘fast’ endocytosis, their size ranged from 2.6 to 3.9 pF (mean = 3.4 ± 0.1 pF; n = 14). Thus, ‘fast’ endocytosis did not appear to occur preferentially in larger cells, although there was a weak correlation between cell size and the amplitude of the ‘fast’ endocytosis (r= 0.47). Nevertheless, the amplitude of the ‘fast’ endocytosis was small (mean = 79 ± 15 fF; n = 13). Given the large variability in the cell size of corticotrophs, we cannot rule out the possibility that ‘fast’ endocytosis may retrieve some of the membrane left over from previous rounds of exocytosis. Unlike ‘slow’ endocytosis which occurred at all [Ca2+]i that stimulated exocytosis, ‘fast’ endocytosis occurred mostly at high [Ca2+]i. In cells stimulated with depolarization pulses, ‘fast’ endocytosis was seen in only 11 % of exo/endocytotic events (n = 37) at [Ca2+]i < 2.5 μm, but increased to 30 % (n = 20) at [Ca2+]i > 3.5 μm. In cells stimulated by flash photolysis of caged Ca2+, none of the exo/endocytotic events (n = 6) showed ‘fast’ endocytosis at [Ca2+]i < 11 μm, but at [Ca2+]i > 13 μm, all exo/endocytotic events (n = 7) showed ‘fast’ endocytosis. ‘Fast’ endocytosis clearly has a higher [Ca2+]i requirement than ‘slow’ endocytosis.

Modulation of endocytosis

Calmodulin has been suggested as the mediator between Ca2+ and endocytosis in bovine chromaffin cells (Artalejo et al. 1996). We therefore tested whether calmodulin also mediated endocytosis in rat corticotrophs. Since Ba2+ is a poor activator of calmodulin (Artalejo et al. 1996; Nucifora & Fox, 1998), we tested whether Ba2+ would support endocytosis in corticotrophs. In this series of experiments, Ca2+ in the bath solution was replaced with Ba2+ (5 mm). A train of depolarization pulses stimulated Ba2+ entry into the cell via voltage-gated Ca2+ channels. Figure 8a shows that exocytosis occurred mostly during the last two depolarization pulses and after the stimulus. The slow exocytosis was not due to intracellular Ca2+ release, because there was no corresponding increase in the fluorescence ratio measured with indo-1 (Fig. 8a). The slow exocytosis was more probably due to the fact that Ba2+ was a poor substitute for Ca2+ in exocytosis and the slow clearance of Ba2+ from the cytosol (Augustine & Eckert, 1984; Przywara et al. 1993). Nevertheless, Fig. 8a shows that both ‘fast’ and ‘slow’ endocytosis were triggered in the presence of Ba2+. In six cells stimulated with Ba2+, three exhibited both ‘fast’ and ‘slow’ endocytosis and the other three showed only ‘slow’ endocytosis. The time constant of ‘slow’ endocytosis was 11.1 ± 0.8 s (mean of 7 exo/endocytotic events from 6 cells), which was about 2-fold larger than that observed for Ca2+-triggered endocytosis (Fig. 3C). The average efficiency of endocytosis was 1.7 ± 0.5 for the seven exo/endocytotic events. Therefore, Ba2+ clearly could support endocytosis, although the kinetics of ‘slow’ endo-cytosis was ≈2-fold slower than in the presence of Ca2+.

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Figure 8. Barium, calmidazolium and cyclosporin A do not inhibit endocytosis

A, Ba2+ can stimulate exocytosis and endocytosis. Ca2+ in the bath solution was replaced with Ba2+ (5 mm) before the cell was stimulated. A train of five 500 ms depolarization pulses delivered at 1.25 Hz stimulated exocytosis which was followed by both ‘fast’ and ‘slow’ endocytosis. The ratio of indo-1 fluorescence only increased slightly with stimulation, suggesting that there was little change in [Ca2+]i. The dotted lines at 0.1 and 1.5 are, respectively, the fluorescence ratios for indo-1 without Ca2+ (Rmin) and saturated with Ca2+ (Rmax). B, the calmodulin inhibitor calmidazolium does not inhibit endocytosis. Endocytosis could be observed after the cell had been dialysed for 5 min with calmidazolium (500 μm) through the whole-cell patch pipette. C, the calcineurin inhibitor cyclosporin A does not inhibit endocytosis. Endocytosis could be observed after the cell was dialysed for 5 min with cyclosporin A (1 μm) through the whole-cell patch pipette.

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To investigate more directly the role of calmodulin in endocytosis in corticotrophs, we dialysed the calmodulin inhibitor calmidazolium (500 μm) into the cell via the whole-cell pipette. Figure 8B shows a cell that was dialysed for about 5 min with calmidazolium and then stimulated with a single 500 ms voltage pulse. The depolarization triggered a [Ca2+]i rise and was accompanied by exocytosis which was followed by endocytosis. For this cell, another voltage pulse given after about 10 min of dialysis also stimulated exocytosis and endocytosis (data not shown). In three cells examined with at least 5 min of dialysis with calmidazolium, one cell exhibited both ‘fast’ and ‘slow’ endocytosis and the other two cells showed only ‘slow’ endocytosis. The time constant of ‘slow’ endocytosis was 7.0 ± 0.8 s (mean of 6 exo/endocytotic events from 3 cells) and the average efficiency of endocytosis was 0.97 ± 0.08. Since endocytosis was observed in the presence of Ba2+ or calmidazolium, calmodulin probably did not mediate endocytosis in rat corticotrophs.

Calcineurin has also been proposed as the Ca2+ sensor for endocytosis (Liu et al. 1994; Lai et al. 1999) and the calcineurin inhibitor cyclosporin A has been reported to modulate endocytosis (Engisch & Nowycky, 1998). We therefore tested whether cyclosporin A (1 μm) could modulate endocytosis in corticotrophs. Corticotrophs were exposed to cyclosporin A for at least 30 min before being patch clamped. A train of depolarization pulses was then delivered in the continued presence of cyclosporin A. In 5 of 5 cells examined, stimulated exocytosis was followed by ‘slow’ endocytosis (Fig. 8C) and ‘fast’ endocytosis was also observed in 2 of the 5 cells. The time constant of ‘slow’ endocytosis in the cyclosporin A-treated cells was 4.8 ± 0.5 s (mean of 6 exo/endocytotic events in 4 cells). Similar results were obtained when cyclosporin A was dialysed into the cell via the whole-cell pipette (5-8 min; n = 5). The average time constant of ‘slow’ endocytosis after at least 5 min of dialysis with cyclosporin A was 6.2 ± 0.6 s and the average efficiency of endocytosis was 1.1 ± 0.1 (mean of 8 exo/endocytotic events from 5 cells). We also tested another calcineurin inhibitor, calcineurin autoinhibitory peptide, which has been shown to modulate endocytosis (Artalejo et al. 1996). In corticotrophs dialysed with calcineurin autoinhibitory peptide (1 mg ml−1) for at least 5 min, stimulated exocytosis was followed by ‘slow’ endocytosis in 4 of 4 cells examined and ‘fast’ endocytosis was observed in one of the cells. The average time constant of ‘slow’ endocytosis in these cells was 6.1 ± 0.7 s and the efficiency of endocytosis was 1.0 ± 0.1 (mean of 7 events from 4 cells). Thus, calcineruin inhibitors have little effect on endocytosis in rat corticotrophs.

Nucleotide dependence of endocytosis

GTP has been reported to be essential for endocytosis in bovine chromaffin cells (Artalejo et al. 1995). In addition, dynamin, a protein that is intimately associated with endocytosis, is a GTPase (van der Bliek, 1999). We therefore tested the GTP dependence of endocytosis in rat corticotrophs. We initially replaced GTP in the pipette solution with the non-hydrolysable GTP analogue GTP-γ-S, but GTP-γ-S stimulated spontaneous exocytosis at resting [Ca2+]i and reduced depolarization-triggered exocytosis. We therefore used another GTP analogue, GDP-β-S, which has also been reported to inhibit endocytosis (Artalejo et al. 1995). In corticotrophs, we have shown previously that intracellular dialysis of GDP-β-S (2 mm) for 5 min can completely abolish the arginine vasopressin (AVP) or the α-adrenergic receptor coupled G-protein response (Tse & Lee, 1998; Tse & Tse, 1998). Therefore, in this experiment, the GTP in the pipette solution was replaced with GDP-β-S (2 mm) and the cell was dialysed for at least 5 min before it was stimulated with a train of depolarization pulses (Fig. 9a). In 6 of 7 cells, stimulated exocytosis was followed by ‘slow’ endocytosis. ‘Fast’ endocytosis was also observed in 3 of the 6 cells. In the presence of GDP-β-S, the time constant of ‘slow’ endocytosis was 4.3 ± 0.6 s and the average efficiency was 0.86 ± 0.07 (mean of 13 exo/endocytotic events from 6 cells). Thus, GDP-β-S has little effect on endocytosis in corticotrophs.

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Figure 9. Nucleotide dependence of endocytosis

A, GDP-β-S fails to inhibit endocytosis. Two 500 ms depolarization pulses were delivered to a cell that was dialysed with GDP-β-S for 5 min. The depolarization-triggered exocytosis was followed by endocytosis. GTP in the pipette solution was replaced with GDP-β-S (2 mm). B, endocytosis is not dependent on ATP. A train of five 500 ms depolarization pulses was delivered to a cell that was dialysed with ATP-γ-S for 4 min. The ATP in the pipette solution was replaced with ATP-γ-S (2 mm). Note that [Ca2+]i recovery was slower in the presence of ATP-γ-S but endocytosis could still be observed. C, endocytosis is not modulated by cAMP. A train of five 250 ms depolarization pulses was delivered at 2 Hz to a cell that was dialysed with cAMP (500 μm) for 5 min. Robust endocytosis could be observed.

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Another nucleotide, ATP, has also been reported to be essential for endocytosis (Heidelberger, 1999). Furthermore, tyrosine phosphorylation has also been reported to modulate endocytosis (Nucifora & Fox, 1999). We therefore tested the ATP dependence of endocytosis in corticotrophs. We replaced the ATP in our pipette solution with ATP-γ-S, a non-hydrolysable analogue of ATP. In cells dialysed with ATP-γ-S, the resting [Ca2+]i became elevated and the recovery of depolarization-triggered [Ca2+]i elevation was slowed down. In the presence of ATP-γ-S, the average time constant of the decay of the Ca2+ transient was 13.0 ± 2.7 s (n = 5), ≈5-fold slower than that recorded with ATP in the pipette solution (2.3 ± 0.1 s, n = 79). Since Ca2+ homeostasis in pituitary cells is strongly dependent on Ca2+-ATPase (Tse et al. 1994), the disruption of Ca2+ homeostasis by ATP-γ-S suggested that the intracellular ATP level was probably low. Nevertheless, ATP-γ-S had little effect on endocytosis. After at least 4 min of dialysis, stimulated exocytosis was followed by ‘slow’ endocytosis in 5 of 6 cells (Fig. 9B) and no ‘fast’ endocytosis was observed in these cells. In the presence of ATP-γ-S, the time constant of ‘slow’ endocytosis was 4.8 ± 0.8 s and the average efficiency was 1.1 ± 0.2 (mean of 5 exo/endocytotic events from 5 cells). Endocytosis in rat corticotrophs is therefore not strongly ATP dependent.

We also considered the effect of cAMP on endocytosis, because cAMP is a major second messenger in the control of ACTH secretion in corticotrophs (Won & Orth, 1990; Lee & Tse, 1997) and cAMP has been reported to modulate the release of granule contents after exocytosis in lactotrophs (Angleson et al. 1999). In this experiment, cAMP (500 μm) was dialysed into the cell via the whole-cell patch pipette. After at least 4 min of dialysis, a train of depolarization pulses was delivered. In 7 of 8 cells examined, stimulated exocytosis was followed by ‘slow’ endocytosis. ‘Fast’ endocytosis was observed in five cells. In the presence of cAMP, the time constant of ‘slow’ endocytosis was 8.6 ± 0.7 s and the average efficiency was 1.3 ± 0.1 (mean of 12 exo/endocytotic events from 7 cells). For the same batch of cells, recorded without cAMP, the time constant of ‘slow’ endocytosis was 7.9 ± 1.0 s and the average efficiency was 1.2 ± 0.1 (mean of 10 exo/endocytotic events from 4 cells). Therefore, cAMP does not modulate endocytosis in rat corticotrophs.

DISCUSSION

  1. Top of page
  2. Abstract
  3. METHODS
  4. RESULTS
  5. DISCUSSION
  6. Acknowledgements

‘Slow’ and ‘fast’ endocytosis

We have found that in rat corticotrophs, endocytosis is strongly coupled to exocytosis. In this study, the first two or three voltage steps (0.5 s duration) of a train of depolarization typically elevated [Ca2+]i to a peak value of several micromolar. A similar [Ca2+]i elevation has been observed in corticotrophs during AVP or α-adrenergic receptor stimulation where [Ca2+]i rose to several micromolar within 1-2 s (Tse & Lee, 1998; Tse & Tse, 1998). Although the [Ca2+]i elevation induced by AVP or α-adrenergic receptor stimulation was due to intracellular Ca2+ release from IP3-sensitive calcium stores and depolarization-stimulated Ca2+ entry was used in this study, we have previously shown that voltage-gated Ca2+ entry and intracellular Ca2+ release generate a similar spatial Ca2+ gradient near the exocytic sites (Tse & Lee, 2000). Thus the [Ca2+]i near the exocytic sites during a train of depolarizations is comparable to that elicited during AVP or α-adrenergic receptor stimulation. At this moderate [Ca2+]i elevation, the endocytosis following stimulated exocytosis is predominantly the ‘slow’ endocytosis which has a time constant of ≈6 s. Endocytosis with a similar time constant has been described in melanotrophs (4 s; Thomas et al. 1994), chromaffin cells (5-15 s; Smith & Neher, 1997; Engisch & Nowycky, 1998), as well as hippocampal neurons (6 s; Klingauf et al. 1998) and retinal bipolar neurons (2 s; von Gersdorff & Matthews, 1994). Previous studies in pancreatic β cells (Eliasson et al. 1996) and chromaffin cells (Smith & Neher, 1997) have shown that endocytosis rapidly disappears within 20-100 s of whole-cell dialysis. However, we found that endocytosis in corticotrophs persisted even after 15 min of whole-cell dialysis. While there was significant rundown of exocytosis and Ca2+ entry during whole-cell dialysis, neither the efficiency nor the kinetics of endocytosis was affected (Fig. 1C and D). These results show that the maintenance of endocytosis is not dependent on small soluble proteins, and that endocytosis is closely coupled to exocytosis in rat corticotrophs.

For many exo/endocytotic events, ‘slow’ endocytosis retrieved more membrane than was added by exocytosis. This excess membrane retrieval is reflected in an average efficiency of endocytosis that is greater than one (1.2 at [Ca2+]i < 3.0 μm; Fig. 2). It occurred at all [Ca2+]i that stimulated exocytosis, and with low or high amplitudes of exocytosis (Fig. 2B and C). This excess membrane retrieval could be intrinsic to endocytosis. Previous studies with electron microscopy have shown that endocytic vesicles are often attached to the plasma membrane through a 25 nm wide neck (Willingham & Pastan, 1983; Takei et al. 1995). This neck is formed by dynamin, which is activated at high [Ca2+]i (Takei et al. 1995). If this neck consists of plasma membrane, it could contribute to the excess membrane that is retrieved. Assuming the diameter of the endocytic vesicle without the neck is the same as that of the exocytosed granule (200 nm) and the neck is 25 nm wide, the neck would have to be 320 nm long to account for the 20 % excess membrane. Necks as long as 700 nm has been reported (Willingham & Pastan, 1983), so the formation of such necks may account for some of the excess membrane retrieval. In a small percentage of exo/endocytotic events (9/91), an efficiency of endocytosis of 2 or more is observed. It seems unlikely that the formation of a neck alone can account for such large excess membrane retrieval, because the neck would have to be 1600 nm or longer. However, it should be noted that 6 of the 9 events with an efficiency of endocytosis > 2 were stimulated by [Ca2+]i transients larger than 3 μm. It is possible that at high [Ca2+]i, the endocytic machinery also retrieves non-granule membrane directly through the formation of endocytic vesicles. The observation that high [Ca2+]i promotes a larger efficiency of endocytosis may therefore reflect the formation of longer necks that connect the endocytic vesicle to the plasma membrane, as well as the formation of endocytic vesicles from non-granule membrane.

In addition to ‘slow’ endocytosis, ‘fast’ endocytosis also followed exocytosis in some exo/endocytotic events. ‘Fast’ endocytosis is different from ‘slow’ endocytosis, because it is not smooth, but contains step-like decreases in Cm and is typically complete within 2 s (Fig. 7). The size of the step-like Cm decrease can be as large as 75 fF, corresponding to a spherical organelle with 1.5 μm diameter. ‘Fast’ endocytosis probably does not retrieve granule membrane that was exocytosed immediately preceding it, because ‘fast’ endocytosis is always followed by ‘slow’ endocytosis, and the amounts of ‘slow’ endocytosis and exocytosis correlate well with each other. Furthermore, ‘fast’ endocytosis has a higher [Ca2+] requirement (> 10 μm) than both exocytosis and ‘slow’ endocytosis. A similar type of ‘fast’ endocytosis has also been described in rat melanotrophs and bovine chromaffin cells (Neher & Zucker, 1993; Thomas et al. 1994; Smith & Neher, 1997). ‘Fast’ endocytosis has been proposed to retrieve membrane that was added by previous rounds of stimulated exocytosis or spontaneous activities (Thomas et al. 1994; Smith & Neher, 1997). In corticotrophs, there is only a weak correlation between the cell size and the amount of ‘fast’ endocytosis (r= 0.47). However, the amount of ‘fast’ endocytosis in these cells is typically small, so the total added membrane from previous exocytic events might not be very large. Therefore, the possibility that ‘fast’ endocytosis may retrieve membrane that was added by previous exocytic events cannot be ruled out here. Alternatively, ‘fast’ endocytosis may retrieve non-granule membrane that could be involved in the normal turnover of the plasma membrane. As in excess membrane retrieval by ‘slow’ endocytosis, high [Ca2+]i may also promote the retrieval of non-granule membrane through ‘fast’ endocytosis.

Exocytosis and endocytosis

When corticotrophs were stimulated with depolarization pulses, ‘slow’ endocytosis typically followed exocytosis and the amount of ‘slow’ endocytosis correlated well with the amount of exocytosis. Furthermore, no endocytosis occurred if no exocytosis was stimulated. These observations suggest that ‘slow’ endocytosis and exocytosis are closely coupled, such that each exocytosed vesicle leads to an endocytosed vesicle. Such close coupling implies that our Cm measurement is a net result of exocytosis and endocytosis. For our depolarization experiments, the temporal overlap of endocytosis and exocytosis is probably small. In corticotrophs, we have previously shown that exocytosis stimulated via voltage-gated Ca2+ entry was due to the generation of a locally high [Ca2+] near the exocytic sites (Tse & Lee, 2000). Thus, following the termination of the train of depolarizations, the spatial Ca2+ gradient would dissipate quickly and little exocytosis would be expected. Consistent with this, simultaneous measurements of amperometry and capacitance in another neuroendocrine cell type, chromaffin cells, have shown that most exocytosis occurred during the depolarization pulses (Engisch & Nowycky, 1998). From our flash photolysis experiments, we find that exocytosis is more steeply Ca2+ dependent than ‘slow’ endocytosis, and that exocytosis is typically a much faster process than ‘slow’ endocytosis (Fig. 4 and Fig. 5). Nevertheless, at low [Ca2+]i or low rates of exocytosis in our flash photolysis experiments, some exocytosis may overlap with endocytosis. Under this condition, we may have overestimated the time constant of ‘slow’ endocytosis and the Ca2+ dependence of ‘slow’ endocytosis may be less steep than that shown in Fig. 5B. However, this does not affect our conclusion that ‘slow’ endocytosis is mediated via a high-affinity Ca2+ sensor (discussed later).

The temporal overlap of exocytosis and endocytosis would affect most strongly the measurement of exocytosis during low [Ca2+]i elevation in our flash photolysis experiments. Figure 4 shows that at lower [Ca2+]i, exocytosis appears to stop, even though [Ca2+]i is still elevated and the readily releasable pool of granules has not been exhausted. This phenomenon can be explained by a close coupling of endocytosis to exocytosis, such that exocytosis and endocytosis of individual vesicles are occurring consecutively. Simulation of a kinetic model with coupled exocytosis and endocytosis shows that cumulative exocytosis can be expected to increase with faster rates of exocytosis and that the time to the peak of exocytosis will also shorten with faster rates of exocytosis (Fig. 6a and B). This model is similar to the ‘kiss-and-run’ model that has been proposed for synaptic vesicle recycling (Palfrey & Artalejo, 1998). Like the ‘kiss-and-run’ model, our model assumes that the same exocytosed vesicle is endocytosed. However, unlike the ‘kiss-and-run’ model, endocytosis in corticotrophs is slow (time constant of 3-6 s). The slow kinetics implies that the exocytosed vesicle in corticotrophs is exposed extracellularly for several seconds. Since the dissolution of the dense core in prolactin-containing granules requires ≈3 s (Angleson et al. 1999), this time interval may allow the dissolution of the dense core in the ACTH-containing granules and thus a more complete release of ACTH. Nevertheless, this time interval is still brief for the mixing of granule and plasma membrane proteins (Thomas et al. 1994). The close coupling between endocytosis and exocytosis therefore not only ensures efficient retrieval of vesicle membrane but also preserves the integrity of the vesicle and plasma membrane.

Ca2+ dependence of endocytosis

Endocytosis has been reported to be both [Ca2+]i independent (Ryan et al. 1996; Wu & Betz, 1996) and [Ca2+]i dependent (Artalejo et al. 1995; Smith & Neher, 1997; Engisch & Nowycky, 1998). In nerve terminals, high [Ca2+]i has been reported to inhibit endocytosis (von Gersdorff & Matthews, 1994; Hsu & Jackson, 1996; Cousin & Robinson, 2000). In corticotrophs, high [Ca2+]i clearly does not inhibit endocytosis, as endocytosis occurs even when [Ca2+]i > 10 μm (Fig. 4a and Fig. 5a). In corticotrophs, endocytosis can also occur at resting [Ca2+]i (< 300 nm). In depolarization experiments, the recovery time course of the stimulated [Ca2+]i transient is three times faster than that of endocytosis, so significant endocytosis occurs after [Ca2+]i has returned to resting levels. Endocytosis therefore does not require high [Ca2+]. Nevertheless, we found that τendo decreased by ≈3-fold when [Ca2+]i increased from 2 to 14 μm (Fig. 3C and Fig. 5B), suggesting that ‘slow’ endocytosis is dependent on Ca2+. Furthermore, the Ca2+ dependence of endocytosis is also suggested by the Ba2+ experiment which shows that substitution of Ba2+ for Ca2+ slows down the kinetics of ‘slow’ endocytosis ≈2-fold (Fig. 8a). Our observation that high [Ca2+]i promotes a larger efficiency of endocytosis (Fig. 2B) also supports the suggestion that endocytosis is Ca2+ dependent. The above observations are consistent with the notion that endocytosis may involve a Ca2+ sensor with a high Ca2+ affinity (Cousin & Robinson, 2000).

It has been proposed that the Ca2+ sensor for endocytosis has a high Ca2+ affinity - a few hundred nanomolar (Lai et al. 1999; Cousin & Robinson, 2000). For endocytosis with such a high Ca2+ affinity, endocytosis would appear to be relatively [Ca2+]i independent at the high level of [Ca2+]i (in the micromolar range) that triggers exocytosis, because the Ca2+ sensor for endocytosis would be saturated at this high [Ca2+]i. A high Ca2+ affinity for endocytosis will also allow endocytosis to occur at resting [Ca2+]i (< 300 nm). Thus our experimental results would be consistent with a high Ca2+ affinity for endocytosis. Direct testing of this proposal will involve examining exo/endocytosis at lower [Ca2+]i (< 1 μm). Unfortunately, in corticotrophs, significant exocytosis could be detected only when depolarization raised [Ca2+]i to > 1 μm (Tse & Lee, 2000). Therefore, in this study, we numerically simulated endocytosis (see Methods) with an EC50 for [Ca2+]i set at 500 nm for a [Ca2+]i transient with a decay time constant of 2.3 s and peak [Ca2+]i of 2 μm. With these parameters, our simulation generated an endocytosis with a time constant of 6.0 s, which is similar to the measured value of 6.4 s at 1.9 μm[Ca2+]i (Fig. 3C) in our depolarization experiments. For a peak [Ca2+]i of 5 μm, the simulation generated an endocytosis with a time constant of 4.2 s which is also close to the measured value of 5.2 s at 5.0 μm (Fig. 3C) in our depolarization experiments. We have suggested earlier in the Discussion that there is little temporal overlap between exocytosis and endocytosis in our depolarization experiments. If there is indeed some temporal overlap, the time constant of endocytosis will be smaller and a Ca2+ sensor of higher affinity (< 500 nm) will be needed to simulate the time course of endocytosis. Nevertheless, endocytosis with a Ca2+ sensor of high Ca2+ affinity could account for the small decrease in the time constant of endocytosis with increasing [Ca2+]i that we measured in our depolarization experiments (Fig. 3C). As for our flash photolysis experiments, the range of [Ca2+]i (8-15 μm) in those experiments would almost completely saturate the Ca2+ sensor for endocytosis, so the time constant of endocytosis should only decrease slightly with increasing [Ca2+]i (Fig. 5B). Therefore, endocytosis in corticotrophs is not [Ca2+]i independent, but instead involves a Ca2+ sensor with high Ca2+ affinity.

The calmodulin-calcineurin-dynamin cascade has been proposed as the mediator between Ca2+ and endocytosis (Liu et al. 1994; Artalejo et al. 1996; Cousin & Robinson 2000). Ca2+ activates calmodulin which stimulates calcineurin. Calcineurin then dephosphorylates dynamin which leads to the formation of endocytic vesicles. We investigated the role of calmodulin in this cascade by substituting Ba2+, a poor activator of calmodulin, for Ca2+ and by intracellular dialysis of calmidazolium, a calmodulin inhibitor. Neither of these manipulations inhibited endocytosis. Ba2+ could clearly support endocytosis, although endocytosis became slower, while calmidazolium had no significant effects on endocytosis (Fig. 8a and B). An alternative to calmodulin as the Ca2+ sensor is the Ca2+-dependent interaction between calcineurin and dynamin (Lai et al. 1999). The EC50 of 0.1-0.4 μm of this interaction is consistent with a high Ca2+ affinity. This interaction is not dependent on the catalytic activity of calcineurin, which is also consistent with our observation that endocytosis is not affected by the calcineurin inhibitors cyclosporin A or calcineurin autoinhibitory peptide. However, this interaction is thought to bring calcineurin into proximity with dynamin and other proteins in the endocytic complex, where it and possibly other phosphatases can dephosphorylate these proteins and thus maintain a functional endocytic complex.

The focus of the Ca2+ activation cascade is dynamin, because dynamin is necessary for endocytosis in neuronal and non-neuronal cells (van der Bliek, 1999). Dynamin is a GTPase that can self-assemble into a ring-like structure which forms the narrow neck that connects an endocytic vesicle to the plasma membrane (Hinshaw & Schmid, 1995; Takei et al. 1995). Dynamin has also been proposed to directly detach the endocytic vesicle from the plasma membrane or to coordinate another molecule for the same function (Sever et al. 1999). Dynamin is activated by dephosphorylation which might belie the reason why substitution of ATP with ATP-γ-S in our pipette solution did not have a significant effect on endocytosis (van der Bliek, 1999). However, substitution of GTP with GDP-β-S in our pipette solution also failed to disrupt endocytosis in corticotrophs. In contrast, under similar experimental conditions, GDP-β-S completely abolished the activation of AVP or the α-adrenergic receptor-coupled G-protein (Tse & Lee, 1998; Tse & Tse, 1998). Since dynamin-GTP is the active form of dynamin, it is possible that GDP-β-S cannot easily displace GTP that is already bound to dynamin. Furthermore, the dynamin GTPase activity is only enhanced during endocytosis, so additional GTP may not be required until endocytosis has exhausted the pool of dynamin-GTP (Sever et al. 1999).

In summary, we have found that endocytosis in corticotrophs is closely coupled temporally and probably spatially to exocytosis. This close coupling is reflected in the good correlation between endocytosis and exocytosis and the persistence of this correlation even after prolonged whole-cell dialysis. Our experimental results are compatible with a slow ‘kiss-and-run’-like model wherein each exocytosed vesicle is eventually endocytosed. We also found that endocytosis involves a Ca2+ sensor with high Ca2+ affinity (EC50≈500 nm), which explains the weak [Ca2+]i dependence at [Ca2+]i levels that stimulate exocytosis, and the continuation of endocytosis at resting [Ca2+]i. With these characteristics for endocytosis, a small [Ca2+]i elevation in rat corticotrophs such as that occurring during CRH stimulation would not only stimulate exocytosis but also ensure efficient endocytosis and thus the sustained release of ACTH.

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Acknowledgements

  1. Top of page
  2. Abstract
  3. METHODS
  4. RESULTS
  5. DISCUSSION
  6. Acknowledgements

We thank Drs Robert J. Kemppainen for the ACTH antibodies, Graham Ellis-Davis for NP-EGTA, and Frederick W. Tse for comments on the manuscript. This work is supported by grants from the Canadian Institutes of Health Research (CIHR) and the Alberta Heritage Foundation for Medical Research (AHFMR). Amy Tse is an AHFMR Senior Scholar.