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. 2000 Feb 15;523(Pt 1):101–115. doi: 10.1111/j.1469-7793.2000.00101.x

On the discrepancy between whole-cell and membrane patch mechanosensitivity in Xenopus oocytes

Yong Zhang 1, Owen P Hamill 1
PMCID: PMC2269787  PMID: 10673547

Abstract

  1. Mechanical stimulation of voltage-clamped Xenopus oocytes by inflation, aspiration, or local indentation failed to activate an increase in membrane conductance up to the point of causing visible oocyte damage.

  2. The absence of mechanosensitivity is not due to the vitelline membrane, rapid MG channel adaptation or tension-sensitive recruitment of new membrane.

  3. Membrane capacitance measurements indicate that the oocyte surface area is at least 5 times larger than that predicted assuming a smooth sphere. We propose that this excess membrane area provides an immediate reserve that can ‘buffer’ membrane tension changes and thus prevent MG channel activation.

  4. High-resolution images of tightly sealed patches and patch capacitance measurements indicate a smooth membrane that is pulled flat and perpendicular across the inside of the pipette. Brief steps of pressure or suction cause rapid and reversible membrane flexing and MG channel activation.

  5. We propose that changes in membrane geometry induced during cell growth and differentiation or as a consequence of specific physiological and pathological conditions may alter mechanosensitivity of a cell independently of the intrinsic properties of channel proteins.

Early after the development of tight seal patch-clamp recording (Sakmann & Neher, 1983), a new class of channel sensitive to mechanical stimulation was identified (Guharay & Sachs, 1984; Methfessel et al. 1986; Morris & Sigurdson, 1989). Subsequently, mechanically gated (MG) channels were reported in a wide variety of cell types (Morris, 1990; Martinac, 1993; Sackin, 1995; Hamill & McBride, 1996; Sachs & Morris, 1998) and based on their ubiquitous expression, it was suggested that they may serve in general cellular functions such as cell volume and cell growth regulation (Taglietti & Toselli, 1988; Sachs, 1988). However in 1991, a study of snail neurons indicated that despite the consistent ability to activate MG K+ channel currents in membrane patches, it was not possible to mechanically activate whole-cell K+ currents (Morris & Horn, 1991). Because this discrepancy cast doubt on the functional role of single MG channels, there was substantial interest in determining whether the discrepancy generalized to other cell types. Subsequent studies of a variety of other cells indicated that whole-cell currents could be mechanically activated (Gustin, 1991; Zhou et al. 1991; Zhou & Kung, 1992; Davis et al. 1992; Cemerikic & Sackin, 1993; Wellner & Isenberg, 1994; Cui et al. 1995; Hu & Sachs, 1996; Setoguchi et al. 1997).

Since the above studies indicate that the discrepancy may not be generalized to all cell types, the question then arises as to nature of the cell-type specific mechanism that might determine its occurrence. In the specific case of snail neurons, it has been suggested that an intact cortical cytoskeleton normally protects the lipid bilayer from undergoing a significant tension increase during mechanical perturbations. In comparison, tight seal formation by disrupting the cortical cytoskeleton causes the membrane patch to become hyper-mechanosensitive (Small & Morris, 1994; Wan et al. 1999; see also Hamill & McBride, 1997). Consistent with this idea was the observation that ‘gently’ sealed patches (see Hamill & McBride, 1992) displayed less mechano-sensitivity than ‘forcefully’ sealed or mechanically traumatized patches (Small & Morris, 1994; Wan et al. 1999; see also Glogauer et al. 1998). Following the same argument, one would predict that cells expressing whole-cell MG currents would not express or retain the same degree of cytoskeleton protection of the bilayer.

Although the above hypothesis is attractive, there are additional observations that complicate this interpretation. In particular, in Xenopus oocytes the high mechano-sensitivity of single MG cation channels is best preserved by gentle sealing protocols and minimal mechanical stimulation of the patch, whereas forceful sealing or mechanical trauma makes the patch hypo-mechanosensitive (Hamill & McBride, 1992, 1997). In this case, it was hypothesized that specific membrane-cytoskeleton interactions were required for optimal mechanosensitivity of the channel. Furthermore, one might predict that the oocyte would be more mechano-sensitive than the patch since membrane-cytoskeleton interactions are relatively undisturbed by whole-cell recording. To directly test this idea and gain further insight into the relationship between patch and whole-oocyte mechanosensitivity, we have measured the response of oocytes to a variety of direct mechanical perturbations. In addition, for completeness, we have tested the mechano-sensitivity of the activated endogenous conductances described in the previous study (Zhang & Hamill, 2000).

METHODS

Xenopus laevis were anaesthetized and oocytes removed. Recovery of the frogs was monitored. Not less than 3 months later a second collection was made after which the anaesthetized frogs were decapitated. See Zhang & Hamill (2000) for full details. The care of frogs and preparation of oocytes, the solutions, chemicals, voltage-clamp, data analysis and patch-clamp recording techniques were the same as described in the preceding paper (Zhang & Hamill, 2000).

In vitro oocyte maturation

To induce oocytes to mature in vitro, immature stage VI oocytes were placed in normal Ringer solution supplemented with 10 μM progesterone for 12 h. Maturation was judged by germinal vesicle breakdown, as indicated by the development of a white spot on the animal pole.

Recording oocyte changes in response to mechanical stimulation

The morphology of oocytes during direct mechanical stimulation was continuously monitored with a video camera attached to the dissection microscope. The images were recorded with a SVHS video recorder for latter capture and display.

Methods of mechanically stimulating whole oocytes

A variety of techniques were used to mechanically stimulate oocytes during simultaneous voltage-clamp and video recording.

(i) Oocyte inflation

To inflate the oocyte, an additional pipette (10–15 μm in tip diameter) was inserted for microinjection of 50 or 100 nl of potassium glutamate solution (150 mm potassium glutamate, free Hepes, pH 7.2), or potassium acetate solution (100 mm potassium acetate, free Hepes, pH 7.2) or 150 KCl solution (150 mm KCl, 5 mm Hepes-KOH, pH 7.2) using a Model NA-1 microinjector (Sutter Instruments). The pH of the injection solutions was adjusted with free Hepes to 7.2.

(ii) Oocyte aspiration

To aspirate the oocyte, a macropipette with a heat-polished tip (30–300 μm diameter) was gently pushed against the oocyte surface and suction (5–15 mmHg) was then applied to aspirate a portion of the oocyte (as much as 10 % volume) into the pipette. Only oocytes with their vitelline membrane intact could be studied by the aspiration method because attempts to aspirate devitellinated oocytes always resulted in oocyte damage. Based on reversible movements of the oocyte in and out of the aspiration pipette, there was little adherence between the vitelline enclosed oocyte and the pipette

(iii) Oocyte localized compression or indentation

Local compression or indentation of the oocyte surface was achieved using either a fluid jet or a heat-polished glass probe. In the first case, a pipette with a tip diameter of ∼15 μm was filled with bath solution and placed ∼10 μm away from the surface of the oocyte. Fluid was ejected toward the oocyte by applying a positive pressure to the pipette causing a visible indentation of the oocyte. In the second case, the oocyte was prodded with a heat sealed pipette mounted on a calibrated micromanipulator (Narishigi model 2101). With these methods, precaution was taken not to apply the stimuli directly to membrane regions near the intracellular electrodes, so as to avoid disruption of the membrane-electrode seals and possible mechano-electrical artifacts.

Capacitance measurements of Xenopus oocytes

The membrane capacitance of oocytes was determined from the transient current response to a step change in command voltage. The membrane current in response to a square wave voltage step is described by eqns (1), (2) and (3).

graphic file with name tjp0523-0101-m1.jpg (1)
graphic file with name tjp0523-0101-m2.jpg (2)
graphic file with name tjp0523-0101-m3.jpg (3)

where I(t) is the membrane current at t. Is and I0 are the steady-state current, and the current before the testing pulse, respectively. Rm, Rs and Cm represent the membrane resistance, electrode resistance and membrane capacitance, respectively. ΔV is the command voltage.

Cm was measured during oocyte inflation. The oocytes were held at −50 mV and a series of voltage pulses of 10 or 5 mV with a 10 ms duration and inter-pulse interval of 10 ms were applied on top of the holding potential. The current response to each voltage pulse was fitted with eqns (1) to (3) and the membrane capacitance and resistance measured from 20 pulses was averaged each second using a program written by Dr Don McBride (University of Texas Medical Branch, TX, USA).

As a positive control for capacitance increase caused by membrane recruitment, the calcium ionophore, A23187 (Sigma), was added to the bath solution (10 μM final concentration) in order to elevate intracellular Ca2+ in progesterone-matured oocytes (see ‘In vitro oocyte maturation’). This procedure triggers a massive fusion of cortical granules with the cell membrane with an accompanying membrane capacitance increase in progesterone-matured oocytes but not in non-matured oocytes (Peres & Bernardini, 1985),

High resolution imaging of cell-attached membrane patches

To improve membrane patch images, pipettes with larger internal tip diameters were used (∼4 μm compared with the standard ∼2 μm; see Zhang & Hamill, 2000). The pipettes were bent using a microforge so that the first 2–3 mm of the tip was parallel to the bottom of the dish (Sokabe & Sachs, 1990) and were not coated with Sylgard. Cell-attached membrane patches were visualized using a long working distance 100 × oil immersion objective lens (Zeiss). The patch images were recorded at 60 frames s−1 using a digital high-speed video camera (Pulnix), triggered by the same computer that controlled the patch and pressure clamp, and captured on line with another computer using a program from Speed Visions (San Diego). Single frames were processed and prepared for publication using Adobe Photoshop (Adobe Systems, Inc). In separate experiments in which the membrane patch area (i.e. in pipettes of similar tip size) was estimated by capacitance measurements (see Methods in Zhang & Hamill, 2000), the pipettes were Sylgard coated to within 100 μm of the tip in order to reduce pipette capacitance and were not bent for high-resolution patch imaging. In all cases, measurements were only made on ‘gently’ sealed patches (see, Hamill & McBride, 1992, 1997).

RESULTS

Oocyte inflation

Figure 1 (top panel) shows the inflation of a voltage-clamped oocyte in response to repeated 50 nl micro-injections of 150 mm potassium glutamate solution. The image taken ∼4 s after the 4th microinjection indicates obvious oocyte inflation. By the 8th microinjection the oocyte had increased to 125 % of its original volume (compared with image 0 before microinjection) but with the 9th injection there was a sudden release of cytoplasmic material from around the injection pipette and a subsequent decrease in cell volume (image R). A stage VI oocyte has a volume of ∼500 nl, so each injection should have increased the volume by ∼10 %. However, the presence of the vitelline membrane may limit expansion of the oocyte (see section below on devitellinated oocytes). The majority of oocytes inflated according to this protocol (28 out of 31 oocytes from 4 frogs) showed no increase in conductance up until the point of rupture when there was a sudden and irreversible increase in current (Fig. 1, bottom panel; see also Fig. 8A). In three oocytes (∼10 %) occasional conductance spikes were observed which seemed to correlate with the microinjections (Fig. 2A). However, similar spikes were also seen in 2 out of 12 oocytes bathed in normal Ringer solution (NR; Zhang & Hamill, 2000) with 20 μM Gd3+ (Fig. 2B). Figure 2C shows a statistical analysis indicating that there was no difference in the occurrence of the spikes in the two solutions. Because of this lack of Gd3+ sensitivity, the spikes are unlikely to be mediated by MG channel activation. Instead, they may be related to the mechanically induced Cl− current spikes that can be induced during Ca2+-activated Cl− current oscillations (Hulsmann et al. 1998).

Figure 1. Response of oocytes the inflation by microinjection.

Figure 1

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Top panel, video images taken before (0), during (4 and 8, images taken ≈4 s after the 4th and 8th injections) and after (R, image taken after the oocyte ruptured) inflation by microinjection; 50 nl of 120 mm potassium glutamate (pH 7.2, adjusted with Hepes) was injected each time. Each unit on the scale bar is 0.04 mm. The inserted voltage electrode (VE), current electrode (IE) and injection pipette (IP) are indicated on the first image. Bottom panel, membrane current response during oocyte inflation. The upper trace shows the membrane current of a voltage-clamped oocyte held at −100 mV. The lower trace indicates when the microinjections were made. At the end of the recording, the membrane current abruptly increased, corresponding to visible damage of the oocyte in the region around the injection pipette.

Figure 8. Membrane conductance and capacitance changes during oocyte inflation.

Figure 8

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A, recording of membrane conductance and capacitance of the same oocyte imaged during inflation in Fig. 1. Upper trace, the volume of the oocyte as a percentage of its volume before any microinjection. Continuous and dashed lines in the middle trace represent the membrane conductance and capacitance. Lower trace, timings of microinjections. B, membrane capacitance and conductance of a progesterone-matured oocyte in response to exposure to 10 μM of the calcium ionophore A23187. Upper trace indicates when the ionophore was applied. NR (10 μl) was applied before ionophore application as a control for mechanical stimulation and showed no effect. Lower panel, changes in membrane capacitance (continuous line) and conductance (dashed line).

Figure 2. Occasional conductance spikes during cell inflation in NR and NR + 20 μM Gd3+.

Figure 2

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A, recording of an oocyte in NR shows a conductance spike induced by the second injection. B, recording of an oocyte in NR with 20 μM Gd2+ shows a conductance spike before any injection and a conductance spike that may have been induced by the first injection. The lower trace in each panel indicates the timing of the injections. C, comparison of the conductance spikes observed in oocytes in NR and NR plus 20 μM Gd3+. Open and hatched bars summarize percentage occurrence of conductance spikes in oocytes in NR (11 oocytes, 2 donor frogs) and NR with 20 μM Gd3+ (12 oocytes, 2 donor frogs), respectively, before any injection (Before), in response to first injection (1st) and versus total number of injections (Total); 100 % would indicate the all oocytes showed conductance spikes before, in response to the first injection and in response to all injections, respectively. Error bars indicate s.e.m.s.

Oocyte aspiration

In an attempt to approximate the mechanical stimulation applied to the membrane in the patch pipette, suction was applied to the whole oocyte using heat-polished macro-pipettes (∼30–300 μm tip diameter). This could result in aspiration of a significant portion (∼10 %) of the oocyte into the pipette (Fig. 3A and B). However, upon removal of the suction, the oocyte rapidly moved out of the pipette. As shown in Fig. 3C, there was no current response to repeated pulses of suction which caused the oocyte to move progressively in and out of the pipette (15 oocytes, 2 frogs, −50 mV in NR). With extreme suction the recordings were lost, possibly due to displacement of electrodes. In these aspiration experiments, it was necessary to leave the vitelline membrane intact to avoid damaging the plasma membrane. As a consequence no tight seal formed between the oocyte and the pipette. Under these circumstances, any membrane tension that develops during aspiration should be isotropic throughout the cell (see Evans, 1989). In contrast, with tight seal formation the tension changes will be mainly restricted to the ‘free’ membrane patch that spans the inside of the pipette (see Fig. 9).

Figure 3. Pipette aspiration of the oocyte does not activate a membrane conductance.

Figure 3

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A and B, images of an oocyte taken before (A) and after (B) aspiration. The oocyte was voltage clamped at −50 mV. C, whole-cell current response to the aspirations. The upper panel indicates the timings of each aspiration. The long pulse indicates a prolonged aspiration. Towards the end of the recording, the aspiration was increased to the point that the voltage clamp was lost.

Figure 9. High resolution video images of a membrane patch and membrane currents during brief steps of pressure and suction.

Figure 9

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A and B, video images of a membrane patch before (0 ms) during (50 ms) and after (250 ms) steps (100 ms duration) of suction (A) and a pressure (B). C and D, the membrane patch currents recorded in the same patch imaged in A and B in response to the suction and pressure steps. The shutter of the digital video camera was gated by the same computer that controlled the patch and pressure clamp.

Oocyte indentation or compression

In order to cause localized indentation or compression of the oocyte a fluid jet from a pipette was directed at the oocyte surface. Oocytes with the vitelline membrane intact (5 oocytes, 1 frog) and oocytes with the vitelline membrane mechanically removed (3 oocytes, 1 frog) were tested. As shown in Fig. 4A and B, a fluid jet caused localized indentation or compression of the membrane. If this stimulus were able to activate only 1 % of the MG channels in the oocytes, the estimated whole-cell current would be at least 200 nA at −100 mV. However, no membrane current was activated up until the fluid jet caused visible damage to the devitellinated oocyte (Fig. 4C and D). As an alternative method for inducing localized indentation of the oocyte, a heat-polished glass probe was pressed directly against the oocyte. As indicated in Fig. 5, sustained indentation (∼300 μm) of the oocyte with a glass probe failed to increase membrane current. Similar null responses were seen in 10 oocytes from 2 frogs. However, rapid (< 1 s) and deeper (> 300 μm) indentations could produce current transients (100–1000 nA) that took 5–30 s to recover after probe withdrawal. These currents were not blocked by either 100 or 500 μM Gd3+ (12 oocytes, 2 frogs, data not shown). Furthermore, the indentations appeared most effective when they displaced or stretched the membrane region near one or both intracellular electrodes (i.e. this membrane region appeared to whiten). The lack of Gd3+ block, the slow recovery after removal of the probe and the apparent interaction with intracellular electrodes indicates that the currents arise from mechanically induced membrane leak and rapid resealing around the electrodes, rather than from MG channel activation. Indeed, similar Gd3+-insensitive currents coud be activated by lateral displacement (i.e. 50–200 μm) of either intracellular electrode within the oocyte (6 oocytes, 1 frog, data not shown).

Figure 4. A devitellinated oocyte stimulated by fluid jets.

Figure 4

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A and B, video images of a devitellinated oocyte before and during fluid jet stimulation. C, image of the same oocyte after it was ruptured by application of a strong fluid jet. D, membrane current recording during the fluid jet applications. The oocyte was held at −100 mV. The sudden increase in membrane current at the end of the recording corresponds to the cell rupture.

Figure 5. Direct indentation of the Xenopus oocyte does not activate an increase in conductance.

Figure 5

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A and B, images of an oocyte voltage clamped at −100 mV before and during indentation (≈300 μm) with a heat-sealed pipette (≈50 μm in diameter). C, recording of membrane current before and during the maintained indentation shown in Fig. 5B.

The vitelline membrane does not prevent oocyte mechanosensitivity

Most inflation experiments were performed on oocytes with their vitelline membrane intact. However, the vitelline membrane may limit the inflation of oocytes and thereby prevent MG channel activation. To exclude this possibility we also tested devitellinated oocytes. As shown in Fig. 6A, a devitellinated oocyte can be inflated to nearly twice its original diameter without being ruptured. However, despite such drastic changes in cell volume, no conductance increase was activated in the majority of oocytes tested (9 out of 11 oocytes from 1 frog; Fig. 6B). In two oocytes, inward current spikes were occasionally associated with microinflations (see Fig. 2) but they were not blocked by 100 μM Gd3+, ruling out a contribution by MG channels (data not shown). As pointed out above, the removal of the vitelline membrane does not unmask a conductance in response to localized compression by a fluid jet (Fig. 4).

Figure 6. Inflation of a devitellinated oocyte fails to increase membrane conductance.

Figure 6

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A and B, images of a devitellinated oocyte before (A) and after (B) microinjections. The diameter of the oocyte before microinjections was ≈1 mm with a volume of ≈500 nl. C, membrane current recording from the same oocyte. No membrane current response was activated with ≈50 microinjections which should cause an ≈5 fold increase in volume; 50 nl of potassium glutamate (titrated with Hepes to pH 7.2) was injected each time. Injections were performed every 10 s.

MG channel adaptation cannot explain mechanical insensitivity

Failure to detect whole MG channel current during oocyte inflation could be due to fast adaptation of the MG channels at hyperpolarized potentials. However, in membrane patches MG channel activity does not display adaptation at strongly depolarized potentials (i.e. at + 50 mV; see Hamill & McBride, 1992 and Fig. 2B in Zhang & Hamill, 2000). For this reason, we investigated whether oocyte inflation could activate a whole-cell current at depolarized potentials. Oocytes were stepped from −50 to +50 mV in potassium acetate solution (120 mm potassium acetate, 2 mm CaCl2, free Hepes, pH 7.2), and during the development of the outward residual current (Ir) the oocyte was microinflated (see Fig. 7). No increase in conductance above the normal smooth development of Ir was seen in a total of five oocytes (from one frog). This result indicates that the depolarization activated Ir is not increased by mechanical stimulation and confirms our earlier observations that Ir is not mediated by MG channels (Zhang & Hamill, 2000).

Figure 7. Response of a depolarized oocyte to mechanical stimulation.

Figure 7

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Recording of the membrane current (middle trace) of an oocyte voltage clamped at +50 mV (upper trace) during several microinjections (lower trace). The bath and pipette solutions were 120 mm potassium acetate, 2 mm CaCl2, 5 mm Hepes-KOH (pH 7.2); 50 nl of potassium glutamate was injected each time.

Recruitment of new membrane does not underlie mechanical insensitivity

Another possible mechanism that may explain the failure to activate whole MG channel currents is that new membrane may be recruited into the plasma membrane (via cytoplasmic membrane pools) preventing or buffering the development of significant tension in the lipid bilayer (see Zorec & Tester, 1993; Dai & Sheetz, 1995; Homann, 1998; Dai et al. 1998). To specifically test if membrane recruitment was induced by oocyte inflation, we monitored membrane capacitance changes simultaneously with membrane conductance during repetitive microinflations. The resolution of our Cm measurements was around 2 nF which would represent a ∼1 % change in whole-oocyte capacitance (see below). As illustrated in Fig. 8A, no significant (< 1 %) change in Cm was detected in the same oocyte shown in Fig. 1, which was inflated to 125 % of its original volume. If membrane recruitment underlies the ability of the oocyte to undergo this increase in size without increasing membrane tension, a 16 % increase in Cm would be expected. Similar results were observed in 10 oocytes recorded in NR (1.5 % ± 0.6 %).

In order to reassure ourselves that we could detect membrane recruitment when in fact it did occur, we monitored Cm changes during the membrane fusion of cortical granules which can be induced by elevating intracellular Ca2+ in hormone-matured but not in non-matured oocytes (Peres & Bernardini, 1985). Specifically, we measured Cm and conductance changes in response to calcium ionophore A23187 application to progesterone-matured oocytes (13 oocytes, 2 frogs). Figure 8B shows that A23187 initially activates a fast and transient increase in membrane conductance, presumably reflecting the Ca2+-activated Cl− conductance (Peres & Bernardini, 1985). After this conductance increase was nearly over and the cell conductance had recovered to near resting levels there was a slow (∼1 min) increase in Cm (∼50 %) which gradually decayed in the maintained presence of A23187. The kinetics and amplitude of this Cm change are consistent with the previous report (Peres & Bernardini, 1985) and indicate that our method can detect membrane recruitment when it does occur.

Excess membrane area of the oocyte

The above Cm measurements indicate that inflation-induced membrane recruitment is not the mechanism by which the oocyte buffers rapid tension changes in the lipid bilayer. However, it may be that the unfolding of the excess membrane area of the oocyte, in the form of microvilli and membrane folds, serves such a role (Bluemink et al. 1983; Zampighi et al. 1995). For example, the predicted Cm of a 1 mm diameter oocyte with a smooth membrane and a specific Cm of 1 μF cm−2 would be 31.4 nF. However, the average Cm of stage VI non-matured oocytes was 166.4 ± 24.7 nF (11 oocytes, 2 frogs) while that of progesterone-matured stage VI oocytes (see Methods) was 70.2 ± 11.6 nF (8 oocytes, 2 frogs). These values are consistent with previous estimates of Cm in stage VI oocytes (Methfessel et al. 1986) and previous studies reporting that progesterone-induced oocyte maturation results in a reduction in the oocyte membrane area (Brummett & Dumont, 1976; Bluemink et al. 1983). Clearly the membrane areas of matured and immature oocytes are much larger than that predicted for a smooth sphere.

Comparison of membrane patch area from patch geometry and patch capacitance measurements

Figure 9A and B shows high-resolution images of a membrane patch after ‘gentle’ tight seal formation. With no application of suction or pressure (top traces), the patch appears as an optically smooth disc that is pulled flat and perpendicular to the walls of the pipette (see also Sokabe & Sachs, 1990). The measured disc radius of ∼3.5 μm indicates a geometric patch area of ∼40 μm2 for the patch in Fig. 9, while the average area of five patches was 55.0 ± 5.2 μm2 (range 39–64 μm2). Assuming a specific Cm of 1 μF cm−2 one would predict a patch Cm of ∼550 fF. Actual capacitance measurements from nine patches in similar sized pipettes, indicated a patch Cm of 447.8 ± 15.6 fF (range 390–510 fF). Thus if anything, Cm measurements underestimate patch area (specific Cm may be < 1 μF cm−2). However, the patch area estimates show good agreement in comparison with the large discrepancy (5-fold) between geometric and capacitance estimates of total oocyte membrane area.

Membrane patch movements in response to suction and pressure steps

Brief (100 ms) application of suction or pressure steps results in a rapid bowing out (away from the tip; Fig. 9A) or in (Fig. 9B) of the membrane patch. Detection of patch movements during the pressure step was limited to 15 ms by the video rate of the digital camera (60 full frames s−1). Following turn-off of the pressure step, the membrane recovers its original flat configuration with a slower time course (250–500 ms). Accompanying these membrane movements is a fast activation and deactivation of MG channel currents (< 10 ms; Fig. 9C and D; see also McBride & Hamill, 1992).

Testing the mechanosensitivity of other activated conductances

In the previous study (Zhang & Hamill, 2000) we described a number of endogenous oocyte currents activated by removal of extracellular Ca2+ (Ic), depolarization (Id), hyperpolarization (Ih) and hypertonicity (Ishrink). Although we have been unable to demonstrate an increase in resting current with mechanical stimulation, we were interested in determining whether these other currents might display mechanosensitivity. Of the four currents, only Ishink displayed a consistent response to oocyte inflation (42 ± 24 % reduction, 4 out of 4 oocytes, 1 frog). Figure 10 shows the effects of inflation on the preactivated Ishrink in two different oocytes. In both recordings, the first inflation produced a large reduction in Ishrink with little response to subsequent inflations. After the initial reduction, Ishrink either displayed slow, partial recovery (Fig. 10A) or no recovery (Fig. 10B). Ic displayed a small, delayed reduction (< 20 %) in response to inflation in some oocytes (4 out of 6, see Zhang, 1998) whereas Ih (data not presented) and Ir (see Fig. 7) were unaffected by oocyte inflation.

Figure 10. The effect of cell inflation on hypertonicity-induced conductance.

Figure 10

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A and B are recordings from two different oocytes. The oocytes were preshrunk in a high potassium Ringer solution solution (120 mm KCl, 2 mm CaCl2, 260 mm mannitol, 5 Hepes-KOH, pH 7.2). Membrane conductance was measured in this solution by voltage ramp. Only the first microinjection induced a significant reduction in membrane conductance which was maintained or slowly increased during the recordings.

DISCUSSION

Despite our expectations from patch-clamp studies, we were unable to mechanically activate a consistent increase in the resting conductance of oocytes. This was not because the mechanical stimuli were below a critical threshold since stimulus strength was routinely increased to values that caused visible oocyte damage. Neither was it due to rapid adaptation of single MG channel activity because a similar lack of activation occurred in oocytes held at strongly positive potentials (+50 mV) where adaptation is absent (Hamill & McBride, 1992). Experiments in which devitellinated oocytes were mechanically stimulated excluded the possibility that the vitelline membrane prevents the activation of MG channels in the whole oocyte.

Another mechanism that may influence oocyte mechano-sensitivity relates to studies of other cell types which indicate that increased membrane tension promotes recruitment of new membrane by fusion of internal vesicles with the plasma membrane (Zorec & Tester, 1993; Dai & Sheetz, 1995; Homann, 1998; Dai et al. 1998). If fast membrane recruitment occurred in the oocyte, it could prevent or ‘buffer’ the development in the bilayer tension during mechanical stimulation. However, our capacitance measurements indicate no increase in membrane area, even when oocytes were inflated to nearly twice their original diameter.

Another mechanism that may buffer membrane tension is related to the large excess membrane area of the oocyte. Our capacitance measurements indicate that stage VI oocytes have a Cm of 166 nF (see also Methfessel et al. 1986) and other studies have reported Cm values almost twice as large (Zampighi et al. 1995). Therefore, oocytes may have 5–10 times the area needed to enclose their geometric volume (31.4 nF for a 1 mm smooth sphere assuming 1 μF cm−2). This large excess membrane area is also consistent with previous EM studies that indicate extensive membrane folding and microvilli (Bluemink et al. 1983; Zampighi et al. 1995). Freeze fracture analysis of Xenopus oocytes by Zampighi et al. (1995) indicate that membrane folds may double the surface area of the oocyte, while microvilli may further increase the membrane area as much as 5-fold. In this analysis the estimated microvilli density was 6–7 microvilli μm−2, with an average microvillus length of 1.4 μm and diameter of 0.12 μm (Zampighi et al. 1995).

The large excess area of the oocyte may provide an immediate membrane reserve that would explain why nearly doubling the diameter of the oocyte by direct inflation was not associated with an increase in membrane capacitance. According to the above measurements, we would have to increase the volume of the oocyte at least 10-fold in order to completely smooth out the membrane. However, such large oocyte inflations of voltage-clamped oocytes have not proven possible, even after removal of the vitelline membrane.

Because there is no specific label for MG channels (analogous to tetrodotoxin for Na+ channels or α-bungarotoxin for the nicotinic channel) it is not possible to determine the distribution of MG channels on membrane specializations such as microvilli and membrane folds. However, if one assumes the channels are evenly distributed over the entire plasma membrane, then most MG channels would be located on microvilli because they make up 80 % of the surface area. The specific location of MG channels could have significant effects on their mechanosensitivity independent of the intrinsic properties of the channel protein. For example, according to Laplace's Law (P = 2T/r), for a given inflation pressure (P), an MG channel located on a microvillus with a radius of curvature (r) of ∼0.05 μm, would experience 1 × 10−4 of the tension (T) compared with an MG channel located on an unfolded region of membrane with a radius of curvature of 500 μm. Furthermore, if a 10 mmHg pressure is required to activate MG channels in a membrane patch with a radius or curvature of 1 μm (Hamill & McBride, 1992), then a pressure of 200 mmHg would be required to activate the same channel located on a microvillus. However, the application of Laplace's Law assumes a thin membrane shell and may not be strictly applicable here because of the presence of the cytoskeleton which maintains microvilli structure as well as supporting the plasma membrane. Nevertheless, these simple calculations illustrate the effect that membrane geometry may have on mechanosensitivity independently of the intrinsic properties of MG channels.

Differential surface distribution of specific membrane proteins or processes may also explain why some oocyte currents but not others are mechanosensitive in the whole oocyte. For example, Ishrink (and perhaps Ic), although not mediated by MG channels, appears to display some mechanosensitivity in that it was inhibited by mechanical stimulation. In contrast, Ir and Ih appeared insensitive to mechanical stimulation. In the case of Ic, it is interesting that a recent study found that gap-junctional conductance in the supporting cells in Corti's organ could be inhibited by increased membrane tension (Zhao & Santos-Sacchi, 1998). Based on the rapid and reversible nature of this inhibition, we suggest it is due not to mechanical destruction of the gap channels but possibly to direct inhibition of the hemi-gap channels (Zhao & Santos-Sacchi, 1998). The inhibition of the Ishrink by oocyte inflation seems to implicate some mechanical mechanism. However, the identification of the transport process underlying Ishrink and what happens to the oocyte's membrane during shrinkage and reinflation needs to be determined. Finally, if the Gd3+-insensitive conductance spikes we (see Fig. 2) and others (Hulsmann et al. 1998) have observed are mediated by Ca2+-activated Cl− channels it will be interesting to determine why this channel, but not the MG channel, can respond to mechanical stimulation of the whole oocyte.

If the immediate membrane reserve of the oocyte serves to prevent tension changes in the bilayer, then presumably the rapid activation of MG channels in the patch indicates a membrane configuration that allows a rapid increase in bilayer tension. Consistent with this idea are high resolution images of membrane patches that indicate an optically smooth membrane that appears to be pulled flat across the inside of the pipette (Fig. 9; see also Sokabe & Sachs, 1990). Furthermore, brief steps of suction or pressure cause the membrane patch to flex rapidly outward or inward accompanied by even more rapid activation and deactivation of MG channels.

According to EM analysis, the oocyte membrane has six or seven microvilli cm−2 (Zampighi et al. 1995). If the microvilli were retained during tight seal formation then a membrane patch with a geometric area of ∼50 μm2 (see Fig. 9) would have ∼300 microvilli, which would increase the membrane area to 250 μm2. If oocyte membrane folds were also retained, the area would increase to 500 μm2 (Zampighi et al. 1995). However, electrical capacitance measurements confirm a patch area of ∼50 μm2. Other lines of evidence also argue against complex membrane patch geometry. For example, high-voltage EM studies of Xenopus oocyte patches (i.e. in pipette tips) indicated the lipid bilayer as a ‘thin veil’ overlying a cortical cytoskeleton meshwork, with no evidence of microvilli (see Figs 11 and 12 of Ruknudin et al. 1991). While with high-resolution light microscopy, one can on occasion resolve what appears to be a motile, brush border-like surface at the oocyte edge (O. P. Hamill, unpublished observations), the tightly sealed membrane patch always appears optically smooth.

Finally, the patch movements associated with MG channel activation are inconsistent with a large excess membrane area that needs to be smoothed out in order to develop tension. For example, if the flat membrane disc in Fig. 9 was bowed into a hemisphere of the same radius (r = 3.5μm), then the membrane area would be doubled (an increase from 38.5 to 77 μm2). However, the height (h) of the patch ‘dome’ is 0.9 μm, so that the actual surface area (A) is increased to only 41 μm2 (A = π(r2+h2)). This simple result indicates that membrane patch area is increased by less than 10 % during MG channel activation.

If the patch differs so dramatically in geometry from the oocyte membrane, then how does tight sealing produce this difference? It may be that during the sealing process the lipid bilayer is decoupled from the underlying cytoskeleton and dragged into the patch as a lipid bleb where it seals tightly to the pipette walls (see Milton & Caldwell, 1990; Hamill & McBride, 1997). In this case, lipid bilayer reserves drawn from the microvilli and membrane folds may form the membrane patch. This phenomenon would be analogous to the drawing out of thin lipid bilayer tethers from red blood cells and other animal cells (Hochmuth & Evans, 1982; Hochmuth et al. 1996). However, such tethers are typically much thinner in diameter than the patch (0.1 μm vs. 2 μm) and are completely free of cytoskeleton (Hochmuth & Evans, 1982). In contrast, there is good evidence that a plug of cytoskeleton is drawn into the pipette along with the membrane which can be decoupled from the membrane by mechanical stimulation of the patch (Ruknudin et al. 1991; Sokabe & Sachs, 1990; Hamill & McBride, 1992; Zhang et al. 2000).

We propose that there may be various types and stages of membrane-cytoskeleton decoupling. For example, in aspirated red blood cells (without tight seal formation) the fluorescently labelled cortical cytoskeleton displays a steep decrease in density along the aspirated projection (Discher et al. 1994). Interestingly, this density gradient can be maintained for the duration of the deformation (> 30 min) but recovers rapidly (within seconds) with release of suction. It may be that tight seal formation creates a similar ‘reversible’ decrease in cytoskeleton density that tends to smooth out the membrane while retaining membrane-cytoskeleton interactions. However, with subsequent over-stimulation of the patch there is an irreversible decoupling of the membrane from the cytoskeleton that results in membrane bleb formation (Hamill & McBride, 1992; Zhang et al. 2000).

An alternative mechanism to explain the change in patch geometry is that the microvilli are sheared off by the initial suction required to form the tight seal and then the membrane rapidly heals as it is drawn into the pipette and seals on the adjacent glass walls. At least consistent with this mechanism is the observation that vesicles can be seen to be swept from the cell surface during seal formation on chick skeletal muscle (Sokabe & Sachs, 1990). Additional studies using membrane fluorescent dyes will be required to determine if this occurs in the oocyte. However, the interesting implication of this mechanism is that the membrane in the patch may be physically altered from the original oocyte membrane and may even be derived from cytoplasmic membrane stores that have a different protein (channel) makeup.

Conclusions and implications for other cell types

Whole-cell and single-channel recordings of heterologously expressed transmitter- and voltage-gated channels in Xenopus oocytes typically give consistent results (Methfessel et al. 1986). This is clearly not the case for the endogenous MG channel. We propose that at least one factor contributing to the discrepancy is the difference in membrane geometry between the two recording configurations. While the oocyte shows a large excess membrane area for its geometry, the tightly sealed patch displays a simple flat geometry consistent with a minimal membrane area. We propose that these geometrical differences affect the ability to rapidly develop tension in the lipid bilayer and activate MG channels.

One of the important adaptations of animal cells (once they lost the protective cell wall of bacteria and plant cells) is a redundancy in their surface membrane in the form of wrinkles, folds and ruffles. This redundancy acts as an immediate membrane reserve that allows the cell to tolerate large cellular deformations and motions without causing stress on the lipid bilayer. For example, when the normally highly deformable red blood cell is pressurized into a smooth sphere it loses its deformability because the plasma membrane can only be further dilated 2–4 % before rupturing. One might therefore expect that many animal cells are relatively insensitive to mechanical stimulation because of these membrane reserves. Conversely, cells lacking such reserves may be rare or may have additional membrane protection mechanisms because of their increased vulnerability to stretch induced bilayer damage.

Considering specific cell types and their reported whole-cell mechanosensitivity, the soma and growth cone of neurones often appears as highly wrinkled and folded (e.g. see Fejtl et al. 1995). This excess membrane area may contribute to the lack of whole-cell mechanosensitivity of molluscan neurones (Morris & Horn, 1991). Another mechanism involving membrane recruitment (from cytoplasmic stores) may serve to protect the same neurones against slower and more maintained deformations (Dai et al. 1998). For the special case of sphereoplasts formed from E. coli, yeast and fungi, the cells are specifically treated to remove their cell wall and the equivalent of the cytoskeleton. As a consequence they assume a simple spherical geometry and, perhaps not surprisingly, display whole-cell MG channel activity when further inflated (Gustin, 1991; Zhou et al. 1991; Zhou & Kung, 1992; Cui et al. 1995). On the other hand, in order to elicit mechanosensitive responses in smooth muscle myocytes, these cells must be over-inflated to the point of forming whole-cell membrane blebs (Setoguchi et al. 1997; Zhang et al. 2000).

Another cell type in which membrane geometry may be an important factor in determining mechanosensitivity is the kidney proximal tubule cell. In this highly polarized cell, MG cation channels are localized to the apical microvilliated (brush border) surface, while MG K+ channels are localized on the relatively smooth proximal surface (Sackin, 1993). Although a whole-cell MG K+ conductance can be activated by osmotic swelling of the cell, apparently the MG cation channels are not sensitive to this stimulus (Cemerekic & Sackin, 1993). Such differential sensitivity is consistent with the idea that the location of the channels on membrane regions of different geometry may influence their mechano-sensitivity. In this case, it will be interesting to determine the membrane geometry and channel distribution in other cell types that display whole-cell mechanosensitive responses (see references in Introduction)

Finally, our studies have focused entirely on the mechano-sensitivity of the stage VI oocyte. However, during oocyte differentiation and maturation there are dramatic changes in membrane geometry. Specifically, stage I oocytes are devoid of microvilli and display a relatively smooth plasma membrane surface (Dick et al. 1970; Dumont, 1972). Also, hormone-matured oocytes show a significant reduction in microvilli density and membrane capacitance (Bluemink et al. 1983). It remains to be determined whether these developmental changes in surface geometry influence the mechanosensitivity of the MG channel and perhaps unmask a functionally important role for this channel.

Acknowledgments

Our research is supported by grants from the National Institute of Arthritis and Musculoskeletal and Skin Diseases, Grant R01-AR42782 and the Muscular Dystrophy Association.

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