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Mapping Fungal Ion Channel Locations
Fungal Genetics and Biology 24, 69–76 (1998) Article No. FG981047
Mapping Fungal Ion Channel Locations
Roger R. Lew Department of Biology, York University, 4700 Keele Street, Toronto, Ontario M3J 1P3, Canada
Accepted for publication February 23, 1998
Lew, R. R., 1998. Mapping fungal ion channel locations. Fungal Genetics and Biology 24, 69–76. Ion channel mapping techniques are described and the results for two fungal organisms, Saprolegnia ferax and Neurospora crassa, are presented. In these species, two channel types have been characterized, stretchactivated channels exhibiting significant calcium permeability and spontaneous channels having significant potassium permeability. Two distinct analyses of patch clamp data, analysis of channel self-clustering and association between different channel types, and localization along the hyphae, reveal significant differences between the two organisms. S. ferax maintains a tip-high gradient of both channel types which is lost after disruption of the actin cytoskeleton. There is significant self-clustering of the channels, as well as interactions between channel types. N. crassa on the other hand does not maintain tip-high gradients, and clustered distributions are observed only for the stretchactivated channels. In terms of physiological roles, evidence is quite strong that the stretch-activated channels function as a growth sensor in S. ferax, but have an unknown function in N. crassa. In both organisms, the potassium permeable channels presumably function in potassium uptake. The differences between these two organisms may be due, in part, to differences in their normal environment: aquatic versus terrestrial. r 1998 Academic Press
Index Descriptors: mechano-sensitive; stretch-activated; calcium; potassium; ion channels; tip growth; ion channel mapping; fungal; Neurospora crassa; Saprolegnia ferax. During the process of fungal hyphal extension (tip growth), ion channels in the plasma membrane may play a 1087-1845/98 $25.00 Copyright r 1998 by Academic Press All rights of reproduction in any form reserved.
variety of essential roles. Some of these roles could be quite general, such as maintenance of intracellular ion concentrations required for metabolism or homeostatic regulation of the membrane electrical potential at a particular value. Some roles could be specific to the process of cellular expansion, such as the generation and maintenance of the osmotic gradients required to control water movement during the increase in cellular volume. Finally, ion channels may control the rate and direction of cellular extension as components of signal transduction systems. In all of the possibilities noted above, the ion channels may or may not exist at specific locations along the growing hypha. There is clear evidence for the asymmetric location of ion transporters based on differences in extracellular ion currents around the tips of hyphae compared to basal regions (Gow, 1984; McGillviray and Gow, 1987; Takeuchi et al., 1988). In some instances, it has been possible to implicate specific ion fluxes, particularly calcium and protons, as playing key roles during tip growth (McGillviray and Gow, 1987; Takeuchi et al., 1988; Lever et al., 1994). Intracellular measurements of hyphal tips of Neurospora crassa confirm that growing tips have distinct electrical characteristics— lower membrane electrical potentials and resistances— compared to nongrowing tips (Table 1). However, even though ion transport at growing tips is different compared with basal regions, it is not clear what makes it so different. With the patch clamp technique, it is possible to map ion channel locations along the tip to determine whether they are asymmetrically located along the tip, play a role in asymmetric ion currents, and are part of the architectural requirements of tip growth. This review will discuss the current advances which have been made in ion channel mapping, starting with a brief description of the patch clamp method, a review of ion channels in fungal species, and analytical techniques useful
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TABLE 1 Electrical Measurements at Neurospora crassa Hyphal Tips: Comparison of Nongrowing versus Growing Tips
Nongrowing Growing
Membrane potential (mV)
Input resistance (MOhm)
2115 6 45 (n 5 17)a 264 6 56 (n 5 9)a
26.3 6 12.1 (n 5 16)b 11.2 6 9.6 (n 5 8)b
Note. The hyphal tips were impaled within 20 µm of the growing apex. Electrical measurements of the membrane potential and input resistance were made using standard electrophysiological techniques (Levina et al., 1991). Data are shown as means 6 standard deviation (n 5 sample size) (N.N. Levina and R.R. Lew, unpublished results). a Significantly different at P 5 0.02. b Significantly different at P 5 0.006.
for determining the nature of channel distributions and densities in the plasma membrane.
PATCH CLAMP METHOD With the patch clamp technique, it is possible to measure ion channel currents in well-defined patches of membrane. The technique has been described in detail elsewhere (Sakmann and Neher, 1995) and its use in fungal species has been recently reviewed (Garrill and Davies, 1994). In fungal species, the key to measuring ion channel activities at various locations along hyphae is retaining hyphal shape after removal of the cell wall. The cell wall must be removed to allow the tip of the patch micropipette access to the plasma membrane. Commonly, the cell wall is removed by enzymatic digestion using cellulases, pectinases, and/or chitinases. Once the wall is removed, the length of the hypha forms a chain of pearls [a string of protoplasts attached by narrow strands (Garrill et al., 1992; Levina et al., 1994, 1995)], with each pearl (protoplast) representing a specific region of the original hypha (Garrill et al., 1993; Levina et al., 1994, 1995). Recent work suggests an alternative method for patch clamping at specific locations along the hypha which may become useful in future work: laser ablation of a region of cell wall to expose the plasma membrane of the cell (Henriksen et al., 1996; Taylor et al., 1996; Henriksen and Assmann, 1997). Once the micropipette tip is appressed to the plasma membrane, slight suction is applied, causing the formation of a high-resistance seal between the membrane and the
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glass rim of the micropipette. For the usual biophysical characterization of ion channels using patch clamp, a seal greater than 1 gigaohm (‘‘gigaseal’’) is used. However, in ion channel mapping, a seal less than 1 gigaohm (‘‘subgigaseal’’ or ‘‘loose patch clamp’’) is commonly used in animal cells (Roberts, 1987; Johnson and Thompson, 1989; Premack et al., 1989; Roberts et al., 1990; Wang and Thompson, 1992; Karpen et al., 1992) and fungal cells (Garrill et al., 1992, 1993; Levina et al., 1994, 1995). One advantage of subgigaseals is the ability to examine relatively large membrane areas. In addition, it avoids structural artifacts of gigaseal formation (Sokabe and Sachs, 1990; Ruknudin et al., 1991; Hamill and McBride, 1997) which can include ion channel ‘‘activity’’ in the absence of protein (Sachs and Qin, 1993). Once channel activity can be measured, the next step is counting the number of channels present in the patch of membrane. This is not necessarily a trivial problem. Channel openings and closings can be measured as steplike transitions in current. The open state of the channel can be identified by the presence of ‘‘shot-noise’’ (MacDonald, 1962) when the channel is open. It is often assumed that the maximum number of concurrent channel openings is equal to the number of channels in the membrane patch. But this assumption may or may not be true. In the case of fungal ion channel mapping, the channels tend to open in bursts (Garrill et al., 1993), validating the assumption. In other cases, if channels open randomly over time, care must be taken to ensure that the estimate of channel numbers will be correct (Horn, 1991; Chang and Kurokawa, 1995). When the number of ion channels is known, there are a variety of analytical techniques that can be used to characterize the distribution of ion channels in the membrane. The patch clamp method therefore offers the ability to directly measure ion channel activity and to analyze ion channel distributions.
FUNGAL ION CHANNELS Most patch clamp research on ion channels in fungal species has used the budding yeast Saccharomyces cerevisiae, which has both K1 selective channels (Gustin et al., 1986) and nonselective mechano-sensitive ion channels (Gustin et al., 1988) on the plasma membrane. One K1 channel (TOK1) has been cloned and characterized (Ketchum et al., 1995; Zhou et al., 1995; Lesage et al., 1996; Vergani et al., 1997). It is an outward rectifier which may function to maintain a negative membrane potential (Bertl
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Mapping Fungal Ion Channel Locations
et al., 1993). There is also evidence that the plasma membrane may contain a Ca21 channel (CCH1) based on sequence homology and a decreased Ca21 accumulation phenotype in a cch1D mutant (Paidhungat and Garrett, 1997). Patch clamp measurements on the plasma membrane of the fission yeast Schizosaccharomyces pombe have also revealed both cation-selective mechano-sensitive ion channels (Zhou and Kung, 1992) and an outwardrectifying K1 channel (Vacata et al., 1993). Filamentous fungi have not been examined as extensively. Besides the work described here on Saprolegnia ferax and N. crassa, Zhou et al. (1991) reported the presence of a cation-selective mechano-sensitive ion channel in germinating urediospores of the rust fungi Uromyces appendiculatus, although questions have been raised about the source of the membrane being patch clamped (Garrill et al., 1992a). Gadolinium inhibits channel activity and, in vivo, germ tube growth and differentiation, suggesting that the channel may function in sensing leaf topography. In the patch clamp experiments using Uromyces, the distribution of the channels along the germ tube was not examined and would certainly shed additional light on their possible physiological roles.
observed in each patch.) Then the x2 is calculated to assess whether the difference between expected and observed distributions is significant. An example of this type of analysis is shown for S. ferax in Table 2. In S. ferax, two distinct channels types are observed: stretch-activated channels which are permeable to calcium and spontaneous channels which are K1 permeable. The analysis is for the distribution of K1 permeable channels in the absence or presence of the actin-disrupting agent cytochalasin E. Under control conditions, the K1 permeable channels are distributed in clusters in the membrane (P , 0.005). This clustering is mediated by the actin cytoskeleton, since disruption of the cytoskeleton causes the channels to distribute randomly in the membrane (0.1. P .0.05). N. crassa has two basic channel types similar to those found in S. ferax: stretch-activated channels permeable to calcium and spontaneous K1 permeable channels (Levina et al., 1995). A similar analysis of channel distribution in N. crassa shows that the K1 permeable channels are distributed randomly, while the stretch-activated channels have a clustered distribution (Table 3). Additional tests can be used to assess whether there is an association between the two channel types in the membrane. Two examples of tests for association include an index of association and two-way contingency tables (Mueller-Dombois and Ellenberg, 1974).
CHANNEL DISTRIBUTION Fundamentally, ion channels may be distributed in membranes in a variety of ways. They may diffuse freely within the membrane and therefore have a random distribution or they may exhibit varying degrees of clustering due to associations between the ion channels, possibly mediated by cytoskeletal elements. In addition, different types of ion channels may or may not associate with one another. Analysis of channel distributions assesses the degrees of clustering and association. The techniques for this type of analysis have been used for a long time in a totally unrelated field of study, quantitative plant ecology (Goldsmith and Harrison, 1976). Quadrat sampling of vegetation is analogous to patch sampling of ion channels, although the scale is very different. The basic test for a random versus clustered (or contagious) distribution of ion channels is a x2 goodness-of-fit test. The number of observed channels per patch is tabulated and compared to the expected distribution if the channels were distributed randomly. (A Poisson distribution is appropriate for the small number of channels
TABLE 2 x2 Test for Presence of Channel Clusters (a ‘‘Contagious’’ Distribution of Channels) in Saprolegnia ferax Number of channels per patch 0
1
2
3
4
Total
9 6.71
0 3.07
41
8 6.59
2 2.40
55
Controla Observed Expected
4 6.58
8 12.04
20 11.01
Cytochalasin Eb Observed Expected
16 12.84
10 18.68
19 13.59
Note. The data are taken from experiments reported in Levina et al. (1994) and are for spontaneous, K1 permeable channels. The number of observed channels per patch were tabulated and compared to the expected distribution if the channels occurred randomly in the plasma membrane. x2 analysis indicated that the channels did in fact occur in clusters. This nonrandom distribution disappeared after treatment with the actin-disrupting agent cytochalasin E, suggesting that actin elements of the cytoskeleton mediate channel clusters. a x2 13.559; degrees of freedom 3; P , 0.005. b x2 7.333; degrees of freedom 3; 0.1 , P , 0.05.
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Roger R. Lew
TABLE 3 x2 Test for Presence of Channel Clusters (a ‘‘Contagious’’ Distribution of Channels) in Neurospora crassa Numbers of channels per patch 0
1
2
3
4
5
Total
0 —
40
1 3.59
39
Spontaneous K1 permeable channelsa Observed Expected
10 13.31
18 14.65
10 8.06
2 2.95
0 —
Stretch-activated Ca21 permeable channelsb Observed Expected
1 2.21
0 6.34
9 9.10
23 8.71
5 6.26
Note. The data are taken from experiments reported in Levina et al. (1995) and are for spontaneous, K1 permeable channels, and stretchactivated Ca21 permeable channels. The number of observed channels per patch was tabulated and compared to the expected distribution if the channels occurred randomly in the plasma membrane. x2 analysis indicated that the spontaneous K1 permeable channels occur randomly, while the stretch-activated Ca21 permeable channels exist in well-defined aggregates. a x2 5.077; degrees of freedom 2; 0.1 , P , 0.05. b x2 32.707; degrees of freedom 4; P , 0.005.
The index of association (IA)1 has the form IA 5 100 3 c/(a 1 b 1 c), where c is the number of patches containing both channels, a is the number of patches containing only one type of channel, and b is the number of patches containing only the other type of channel. For the two fungal species we have examined, the IA is 58.0% (S. ferax) or 73.7% (N. crassa). The advantage of this technique is that it does not require knowledge of the number of patches lacking either channel or the actual number of channels in a patch. The disadvantage is that there is a possibility that the analysis can be misleading. This becomes clear if the two-way contingency test is used to analysis data from S. ferax. In a two-way contingency test, the observed numbers of patches containing both channel types, one or the other channel type, or no channels are compared to the expected values if the channels were distributed randomly. For S. ferax, a x2test of the observed and expected values reveals a highly significant association between the two channel types, but also a high proportion of K1 permeable channels which are always found alone, without stretch-activated channels (Table 4). The index of association is unable to reveal the presence of these two distinct K1 permeable channel populations. 1
Abbreviation used: IA, index of association.
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The relationship between the two channel types in the plasma membrane is clearly shown for S. ferax and N. crassa in Fig. 1. In S. ferax, there are two spatially distinct populations of K1 permeable channels. One population is found in the absence of stretch-activated channels, while the other population is always associated with stretchactivated channels. The K1 permeable/stretch-activated channel clusters are not disrupted by cytochalasin (Levina et al., 1994). By comparison, two spatially distinct K1 permeable channel populations are absent in N. crassa, and the association between stretch-activated channels and K1 permeable channels is weak, almost statistically nonsignficant. To summarize, analysis of distributions clearly shows that different types of ion channels in fungal plasma membrane exhibit distinct patterns of distribution: random, clustered, and/or associated with one another. At least in the case of S. ferax, nonrandom clustering of either channel type can be attributed to some actin-mediated connection to the cytoskeleton, although clusters of both channel types together must rely upon some other mechanism. The analysis up to now does not directly address asymmetric distributions of ion channels along the fungal hypha. Although x2tests can be used to analyze channel numbers in protoplasts from apical versus basal regions of the hypha (Garrill et al., 1992), the best determinant for asymmetric distribution relies upon direct examination of channel densities.
CHANNEL MAPPING Determining channel densities along a fungal hypha first requires knowledge of the surface area of the patch of
TABLE 4 Two-Way Contingency Test for Association of Spontaneous and Stretch-Activated Channels in Saprolegnia feraxa Spontaneous channels
Stretch-activated channels Present Absent
Present
Absent
40 (17.14) 24 (14.86)
5 (5.36) 15 (4.64)
Note. Data are from Levina et al. (1994). The observed and expected (in parentheses) numbers of observations are shown for presence and absence of channel type. a x2 59.26; degrees of freedom 2; P . 0.005.
Mapping Fungal Ion Channel Locations
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membrane within the micropipette. The most accurate measurements can be performed using scanning electron microscopy, but unfortunately, this is a destructive assay. Instead, we have determined the relationship between bubble number (Mittman et al., 1987) and inside diameters measured from scanning electron micrographs and then used this linear regression to estimate patch area (Levina et al., 1994). The bubble number is a measure of the pressure required to pass air through the micropipette tip into methanol. Measurements of micropipette tip resistance may also be used, but the resistance varies with
FIG. 2. Channel densities along the hypha for Saprolegnia ferax. Stretch-activated channels are shown in the upper panel and spontaneous potassium permeable channels are shown in the lower panel for controls (circles) and cytochalasin E treatment (squares). The distance from the tip was calculated using the mean diameters of protoplasts originating from the hyphal tip, second protoplast from the tip, etc. The tip-high gradient of stretch-activated channels is statistically significant; the peak at the third protoplast from the tip for cytochalasin treatment is not. Data (mean 6 standard error, sample size ranged from 4 to 21) are redrawn from Levina et al. (1994).
FIG. 1. Relationship between stretch-activated (SA) calcium permeable and spontaneous potassium permeable ion channel densities for Saprolegnia ferax (top) and Neurospora crassa (bottom). Data are from Levina et al. (1994, 1995).
salt concentration in bathing and micropipette filling solutions, so errors are very likely. Channel densities can then be determined for apical to basal regions of the hypha. Results for S. ferax are shown in Fig. 2 and for N. crassa in Fig. 3.
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Roger R. Lew
ROLES IN TIP GROWTH
FIG. 3. Channel densities along the hypha for Neurospora crassa. Stretch-activated channels are shown in the upper panel and spontaneous potassium permeable channels are shown in the lower panel. Data (mean 6 standard error, sample size ranged from 3 to 10) are from Levina et al. (1995).
In the oomycete water mold S. ferax. there are tip-high gradients of spontaneous K1 permeable channels and stretch-activated Ca21 permeable channels (both gradients are statistically significant). Interestingly, actin disruption with cytochalasin E causes both tip-high gradients to disappear. This is strong evidence that the actin cytoskeleton controls channel location at the hyphal apex (Fig. 2). In the ascomycete N. crassa, there is no indication of a tip-high gradient of either the K1 permeable channels or stretch-activated channels (Fig. 3).
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Intuitively, if ion channels are explicitly required during the process of tip growth, one would expect them to be localized to the tip-growing region of the hypha. This is true in the case of the aquatic organism S. ferax. Potential roles for these channels in tip growth have been established (Garrill et al., 1993). Because the lanthinide gadolinium inhibits the stretch-activated channels, inhibits growth, and abolishes a tip-high intracellular Ca21 gradient observed during growth, the stretch-activated channels probably act as a growth sensor, That is, during tip expansion, activation of the channels would cause a localized influx of Ca21, which would act as a signal for continued secretion of membrane and cell wall material at the tip, thus maintaining tip expansion. The K1 permeable channels are inhibited by tetraethylammonium, which inhibits tip expansion transiently. Because the K1 movement is into the cell, the K1 permeable channels may function in maintaining intracellular osmolarity during the volume increase due to tip expansion. Because of the transient effect of channel inhibition, it is clear that the cell can utilize other mechanisms for maintaining intracellular osmolarity. In the terrestrial organism N. crassa, the absence of tip-high gradients of either channel type implies that the channels do not have a function in tip expansion. Indeed, the inhibition of the stretch-activated channels or K1 permeable channels with gadolinium or tetraethylammonium only transiently inhibits growth. It is possible that these striking differences between these two fungi are a result of evolution in two very different environments. In air, the fungal hyphal tip does not have an assured supply of extracellular Ca21 to rely upon as a component of a ‘‘growth-sensor’’ signal transduction system, and water required for volume increases may in fact have to travel considerable distances within the hypha to the growing tip. By contrast, evolution in an aquatic environment means that Ca21 and water are directly accessible to the growing tip. Hence, S. ferax would have no difficulty utilizing stretch-activated channels and K1 permeable channels in the tip growth process. Although there has been considerable research on channel location and clustering in animal neuronal cells (for a recent review, see Whatley and Harris, 1996), very few studies have been performed in nonanimal cells. Besides the work cited above using fungal organisms, Taylor et al. (1996) examined mechano-sensitive ion channel location in the fucus rhizoid. Similar to the results with N. crassa, they found no evidence for an asymmetric
Mapping Fungal Ion Channel Locations
localization of mechano-sensitive channels in these highly polarized algal zygotes (based on presence/absence criteria and channel density). In this system, Taylor et al. (1996) suggest that possible physiological functions for these channels could include osmoregulation and cytosolic Ca21 modulation of polar extension. Based upon the results to date, it is clear that there is a wealth of diversity in channel localization in the plasma membrane. Many questions regarding the extent to which this membrane architecture regulates cellular growth and morphological development remain to be explored. For nonanimal cells, the development of the laser ablation technique and its use in ion channel mapping in Fucus extend our knowledge to another kingdom besides the fungi. But within the fungal kingdom, there is a lot of work to be done.
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75 Henriksen, G. H., and Assmann, S. M. 1997. Laser-assisted patchclamping: A methodology. Pflugers Arch. 433: 832–841. Horn, R. 1991. Estimating the number of channels in patch recordings. Biophys. J. 60: 433–439. Johnson, J. W., and Thompson, S. 1989. Measurement of nonuniform current densities and current kinetics in Aplysia neurons using large patch method. Biophys. J. 55: 299–308. Karpen, J. W., Loney, J. W., and Baylor, D. A. 1992. Cyclic GMP-activated channels of salamander retinal rods: Spatial distribution and variation of responsiveness. J. Physiol. 488: 257–274. Ketchum, K. A., Joiner, W. J., Sellers, A. J., Kaczmarek, L. K., and Goldstein, S. A. N. 1995. A new family of outwardly rectifying potassium channel proteins with two pore domains in tandem. Nature 376: 690–695. Lesage, F., Guillemare, E., Fink, M., Duprat, F., Lazdunski, M., Romey, G., and Barhanin, J. 1996. A pH-sensitive yeast outward rectifier K1 channel with two pore domains and novel gating properties. J. Biol. Chem. 271: 4183–4187. Lever, M. C., Robertson, B. E. M., Buchan, A. D. B., Miller, P. F. P., Gooday, G. W., and Gow, N. A. R. 1994. pH and Ca21 dependent galvanotropism of filamentous fungi: Implications and mechanisms. Mycol. Res. 98: 301–306. Levina, N. N., Lew, R. R., Heath, I. B. 1994. Cytoskeletal regulation of ion channel distribution in the tip-growing organism Saprolegnia ferax. J. Cell Sci. 107: 127–134. Levina, N. N., Lew, R. R., Hyde, G. J., and Heath, I. B. 1995. The roles of calcium ions and plasma membrane ion channels in hyphal tip growth of Neurospora crassa. J. Cell Sci. 108: 3405–3417. MacDonald, D. K. C. 1962. Noise and Fluctuations: An Introduction. Wiley, New York/London. McGillviray, A. M., and Gow, N. A. R. 1987. The transhyphal electrical current of Neurospora crassa is carried principally by protons. J. Gen. Microbiol. 133: 2875–2881. Mittman, S., Flaming, D. G., Copenhagen, D. R., and Belgium, J. H. 1987. Bubble pressure measurement of micropipet tip outer diameter. J. Neurosci. Methods 22: 161–165. Mueller-Dombois, D., and Ellenberg, H. 1974. Aims and Methods of Vegetation Ecology. Wiley, New York/Toronto. Paidhungat, M., and Garrett, S. 1997. A homolog of mammalian, voltage-gated calcium channels mediates yeast pheromone-stimulated Ca21 uptake and exacerbates the cdc1(Ts) growth defect. Mol. Cell. Biol. 17: 6339–6347. Premack, B. A., Thompson, S., and Coombs-Hahn, J. 1989. Clustered distribution and variability in kinetics of transient K channels in molluscan neuron cell bodies. J. Neurosci. 9: 4089–4099. Roberts, W. M. 1987. Sodium channels near end-plates and nuclei of snake skeletal muscle. J. Physiol. 388: 213–232. Roberts, W. M., Jacobs, R. A., and Hudspeth, A. J. 1990. Colocalization of ion channels involved in frequency selectivity and synaptic transmission at presynaptic active zones of hair cells. J. Neurosci. 10: 3664–3684. Ruknudin, A., Song, M. J., and Sachs, F. 1991. The ultrastructure of patch-clamped membranes: A study using high voltage electron microscopy. J. Cell Biol. 112: 125–134. Sachs, F., and Qin, F. 1993. Gated, ion-selective channels observed with
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76 patch pipettes in the absence of membranes: Novel properties of a gigaseal. Biophys. J. 65: 1101–1107. Sakmann, B., and Neher, E. 1995. Single-Channel Recording, 2nd ed. Plenum, New York/London. Sokabe, M., and Sachs, F. 1990. The structure and dynamics of patch-clamped membranes: A study using differential interference contrast light microscopy. J. Cell Biol. 111: 599–606. Takeuchi, Y., Schmid, J., Caldwell, J. H., and Harold, F. M. 1988. Transcellular ion currents and extension of Neurospora crassa hyphae. J. Membr. Biol. 101: 33–41. Taylor, A. R., Manison, N. F. H., Fernandez, C., Wood, J., and Brownlee, C. 1996. Spatial organization of calcium signalling involved in cell volume control in the fucus rhizoid. Plant Cell 8: 2015–2031. Vacata, V., Hofer, M., Larsson, P., and Lecar, H. 1993. Ionic channels in the plasma membrane of Schizosaccharomyces pombe: Evidence from patch-clamp experiments. J. Bioenerg. Biomembr. 24: 43–53. Vergani, P., Miosga, T., Jarvis, S. M., and Blatt, M. R. 1997. Extracellular K1 and Ba21 mediate voltage-dependent inactivation of the outward-
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Roger R. Lew
rectifying K1 channel encoded by the yeast gene TOK1. FEBS Lett. 405: 337–344. Wang, S. S., and Thompson, S. 1992. A-type potassium channel clusters revealed using a new statistical analysis of loose patch data. Biophys. J. 63: 1018–1025. Whatley, V. L., and Harris, R. A. 1996. The cytoskeleton and neurotransmitter receptors. Int. Rev. Neurobiol. 39: 113–143. Worden, M. K., Rahamimoff, R., and Kravitz, E. A. 1994. A voltagesensitive cation channel present in clusters in lobster skeletal muscle membrane. J. Membr. Biol. 141: 167–175. Zhou, X-L., Stumpf, M. A., Hoch, H,C., and Kung, C. 1991. A mechanosensitive channel in whole cells and in membrane patches of the fungus Uromyces. Science 253: 1415–1417. Zhou, X-L., and Kung, C. 1992. A mechanosensitive ion channel in Schizosaccharomyces pombe. EMBO J. 11: 2869–2875. Zhou, X-L., Vaillant, B., Loukin, S. H., Kung, C., and Saimi, Y. 1995. YKC1 encodes a depolarization-activated K1 channel in the plasma membrane of yeast. FEBS Lett. 373: 170–176.
Mapping Fungal Ion Channel Locations
Roger R. Lew Department of Biology, York University, 4700 Keele Street, Toronto, Ontario M3J 1P3, Canada
Accepted for publication February 23, 1998
Lew, R. R., 1998. Mapping fungal ion channel locations. Fungal Genetics and Biology 24, 69–76. Ion channel mapping techniques are described and the results for two fungal organisms, Saprolegnia ferax and Neurospora crassa, are presented. In these species, two channel types have been characterized, stretchactivated channels exhibiting significant calcium permeability and spontaneous channels having significant potassium permeability. Two distinct analyses of patch clamp data, analysis of channel self-clustering and association between different channel types, and localization along the hyphae, reveal significant differences between the two organisms. S. ferax maintains a tip-high gradient of both channel types which is lost after disruption of the actin cytoskeleton. There is significant self-clustering of the channels, as well as interactions between channel types. N. crassa on the other hand does not maintain tip-high gradients, and clustered distributions are observed only for the stretchactivated channels. In terms of physiological roles, evidence is quite strong that the stretch-activated channels function as a growth sensor in S. ferax, but have an unknown function in N. crassa. In both organisms, the potassium permeable channels presumably function in potassium uptake. The differences between these two organisms may be due, in part, to differences in their normal environment: aquatic versus terrestrial. r 1998 Academic Press
Index Descriptors: mechano-sensitive; stretch-activated; calcium; potassium; ion channels; tip growth; ion channel mapping; fungal; Neurospora crassa; Saprolegnia ferax. During the process of fungal hyphal extension (tip growth), ion channels in the plasma membrane may play a 1087-1845/98 $25.00 Copyright r 1998 by Academic Press All rights of reproduction in any form reserved.
variety of essential roles. Some of these roles could be quite general, such as maintenance of intracellular ion concentrations required for metabolism or homeostatic regulation of the membrane electrical potential at a particular value. Some roles could be specific to the process of cellular expansion, such as the generation and maintenance of the osmotic gradients required to control water movement during the increase in cellular volume. Finally, ion channels may control the rate and direction of cellular extension as components of signal transduction systems. In all of the possibilities noted above, the ion channels may or may not exist at specific locations along the growing hypha. There is clear evidence for the asymmetric location of ion transporters based on differences in extracellular ion currents around the tips of hyphae compared to basal regions (Gow, 1984; McGillviray and Gow, 1987; Takeuchi et al., 1988). In some instances, it has been possible to implicate specific ion fluxes, particularly calcium and protons, as playing key roles during tip growth (McGillviray and Gow, 1987; Takeuchi et al., 1988; Lever et al., 1994). Intracellular measurements of hyphal tips of Neurospora crassa confirm that growing tips have distinct electrical characteristics— lower membrane electrical potentials and resistances— compared to nongrowing tips (Table 1). However, even though ion transport at growing tips is different compared with basal regions, it is not clear what makes it so different. With the patch clamp technique, it is possible to map ion channel locations along the tip to determine whether they are asymmetrically located along the tip, play a role in asymmetric ion currents, and are part of the architectural requirements of tip growth. This review will discuss the current advances which have been made in ion channel mapping, starting with a brief description of the patch clamp method, a review of ion channels in fungal species, and analytical techniques useful
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TABLE 1 Electrical Measurements at Neurospora crassa Hyphal Tips: Comparison of Nongrowing versus Growing Tips
Nongrowing Growing
Membrane potential (mV)
Input resistance (MOhm)
2115 6 45 (n 5 17)a 264 6 56 (n 5 9)a
26.3 6 12.1 (n 5 16)b 11.2 6 9.6 (n 5 8)b
Note. The hyphal tips were impaled within 20 µm of the growing apex. Electrical measurements of the membrane potential and input resistance were made using standard electrophysiological techniques (Levina et al., 1991). Data are shown as means 6 standard deviation (n 5 sample size) (N.N. Levina and R.R. Lew, unpublished results). a Significantly different at P 5 0.02. b Significantly different at P 5 0.006.
for determining the nature of channel distributions and densities in the plasma membrane.
PATCH CLAMP METHOD With the patch clamp technique, it is possible to measure ion channel currents in well-defined patches of membrane. The technique has been described in detail elsewhere (Sakmann and Neher, 1995) and its use in fungal species has been recently reviewed (Garrill and Davies, 1994). In fungal species, the key to measuring ion channel activities at various locations along hyphae is retaining hyphal shape after removal of the cell wall. The cell wall must be removed to allow the tip of the patch micropipette access to the plasma membrane. Commonly, the cell wall is removed by enzymatic digestion using cellulases, pectinases, and/or chitinases. Once the wall is removed, the length of the hypha forms a chain of pearls [a string of protoplasts attached by narrow strands (Garrill et al., 1992; Levina et al., 1994, 1995)], with each pearl (protoplast) representing a specific region of the original hypha (Garrill et al., 1993; Levina et al., 1994, 1995). Recent work suggests an alternative method for patch clamping at specific locations along the hypha which may become useful in future work: laser ablation of a region of cell wall to expose the plasma membrane of the cell (Henriksen et al., 1996; Taylor et al., 1996; Henriksen and Assmann, 1997). Once the micropipette tip is appressed to the plasma membrane, slight suction is applied, causing the formation of a high-resistance seal between the membrane and the
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glass rim of the micropipette. For the usual biophysical characterization of ion channels using patch clamp, a seal greater than 1 gigaohm (‘‘gigaseal’’) is used. However, in ion channel mapping, a seal less than 1 gigaohm (‘‘subgigaseal’’ or ‘‘loose patch clamp’’) is commonly used in animal cells (Roberts, 1987; Johnson and Thompson, 1989; Premack et al., 1989; Roberts et al., 1990; Wang and Thompson, 1992; Karpen et al., 1992) and fungal cells (Garrill et al., 1992, 1993; Levina et al., 1994, 1995). One advantage of subgigaseals is the ability to examine relatively large membrane areas. In addition, it avoids structural artifacts of gigaseal formation (Sokabe and Sachs, 1990; Ruknudin et al., 1991; Hamill and McBride, 1997) which can include ion channel ‘‘activity’’ in the absence of protein (Sachs and Qin, 1993). Once channel activity can be measured, the next step is counting the number of channels present in the patch of membrane. This is not necessarily a trivial problem. Channel openings and closings can be measured as steplike transitions in current. The open state of the channel can be identified by the presence of ‘‘shot-noise’’ (MacDonald, 1962) when the channel is open. It is often assumed that the maximum number of concurrent channel openings is equal to the number of channels in the membrane patch. But this assumption may or may not be true. In the case of fungal ion channel mapping, the channels tend to open in bursts (Garrill et al., 1993), validating the assumption. In other cases, if channels open randomly over time, care must be taken to ensure that the estimate of channel numbers will be correct (Horn, 1991; Chang and Kurokawa, 1995). When the number of ion channels is known, there are a variety of analytical techniques that can be used to characterize the distribution of ion channels in the membrane. The patch clamp method therefore offers the ability to directly measure ion channel activity and to analyze ion channel distributions.
FUNGAL ION CHANNELS Most patch clamp research on ion channels in fungal species has used the budding yeast Saccharomyces cerevisiae, which has both K1 selective channels (Gustin et al., 1986) and nonselective mechano-sensitive ion channels (Gustin et al., 1988) on the plasma membrane. One K1 channel (TOK1) has been cloned and characterized (Ketchum et al., 1995; Zhou et al., 1995; Lesage et al., 1996; Vergani et al., 1997). It is an outward rectifier which may function to maintain a negative membrane potential (Bertl
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Mapping Fungal Ion Channel Locations
et al., 1993). There is also evidence that the plasma membrane may contain a Ca21 channel (CCH1) based on sequence homology and a decreased Ca21 accumulation phenotype in a cch1D mutant (Paidhungat and Garrett, 1997). Patch clamp measurements on the plasma membrane of the fission yeast Schizosaccharomyces pombe have also revealed both cation-selective mechano-sensitive ion channels (Zhou and Kung, 1992) and an outwardrectifying K1 channel (Vacata et al., 1993). Filamentous fungi have not been examined as extensively. Besides the work described here on Saprolegnia ferax and N. crassa, Zhou et al. (1991) reported the presence of a cation-selective mechano-sensitive ion channel in germinating urediospores of the rust fungi Uromyces appendiculatus, although questions have been raised about the source of the membrane being patch clamped (Garrill et al., 1992a). Gadolinium inhibits channel activity and, in vivo, germ tube growth and differentiation, suggesting that the channel may function in sensing leaf topography. In the patch clamp experiments using Uromyces, the distribution of the channels along the germ tube was not examined and would certainly shed additional light on their possible physiological roles.
observed in each patch.) Then the x2 is calculated to assess whether the difference between expected and observed distributions is significant. An example of this type of analysis is shown for S. ferax in Table 2. In S. ferax, two distinct channels types are observed: stretch-activated channels which are permeable to calcium and spontaneous channels which are K1 permeable. The analysis is for the distribution of K1 permeable channels in the absence or presence of the actin-disrupting agent cytochalasin E. Under control conditions, the K1 permeable channels are distributed in clusters in the membrane (P , 0.005). This clustering is mediated by the actin cytoskeleton, since disruption of the cytoskeleton causes the channels to distribute randomly in the membrane (0.1. P .0.05). N. crassa has two basic channel types similar to those found in S. ferax: stretch-activated channels permeable to calcium and spontaneous K1 permeable channels (Levina et al., 1995). A similar analysis of channel distribution in N. crassa shows that the K1 permeable channels are distributed randomly, while the stretch-activated channels have a clustered distribution (Table 3). Additional tests can be used to assess whether there is an association between the two channel types in the membrane. Two examples of tests for association include an index of association and two-way contingency tables (Mueller-Dombois and Ellenberg, 1974).
CHANNEL DISTRIBUTION Fundamentally, ion channels may be distributed in membranes in a variety of ways. They may diffuse freely within the membrane and therefore have a random distribution or they may exhibit varying degrees of clustering due to associations between the ion channels, possibly mediated by cytoskeletal elements. In addition, different types of ion channels may or may not associate with one another. Analysis of channel distributions assesses the degrees of clustering and association. The techniques for this type of analysis have been used for a long time in a totally unrelated field of study, quantitative plant ecology (Goldsmith and Harrison, 1976). Quadrat sampling of vegetation is analogous to patch sampling of ion channels, although the scale is very different. The basic test for a random versus clustered (or contagious) distribution of ion channels is a x2 goodness-of-fit test. The number of observed channels per patch is tabulated and compared to the expected distribution if the channels were distributed randomly. (A Poisson distribution is appropriate for the small number of channels
TABLE 2 x2 Test for Presence of Channel Clusters (a ‘‘Contagious’’ Distribution of Channels) in Saprolegnia ferax Number of channels per patch 0
1
2
3
4
Total
9 6.71
0 3.07
41
8 6.59
2 2.40
55
Controla Observed Expected
4 6.58
8 12.04
20 11.01
Cytochalasin Eb Observed Expected
16 12.84
10 18.68
19 13.59
Note. The data are taken from experiments reported in Levina et al. (1994) and are for spontaneous, K1 permeable channels. The number of observed channels per patch were tabulated and compared to the expected distribution if the channels occurred randomly in the plasma membrane. x2 analysis indicated that the channels did in fact occur in clusters. This nonrandom distribution disappeared after treatment with the actin-disrupting agent cytochalasin E, suggesting that actin elements of the cytoskeleton mediate channel clusters. a x2 13.559; degrees of freedom 3; P , 0.005. b x2 7.333; degrees of freedom 3; 0.1 , P , 0.05.
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Roger R. Lew
TABLE 3 x2 Test for Presence of Channel Clusters (a ‘‘Contagious’’ Distribution of Channels) in Neurospora crassa Numbers of channels per patch 0
1
2
3
4
5
Total
0 —
40
1 3.59
39
Spontaneous K1 permeable channelsa Observed Expected
10 13.31
18 14.65
10 8.06
2 2.95
0 —
Stretch-activated Ca21 permeable channelsb Observed Expected
1 2.21
0 6.34
9 9.10
23 8.71
5 6.26
Note. The data are taken from experiments reported in Levina et al. (1995) and are for spontaneous, K1 permeable channels, and stretchactivated Ca21 permeable channels. The number of observed channels per patch was tabulated and compared to the expected distribution if the channels occurred randomly in the plasma membrane. x2 analysis indicated that the spontaneous K1 permeable channels occur randomly, while the stretch-activated Ca21 permeable channels exist in well-defined aggregates. a x2 5.077; degrees of freedom 2; 0.1 , P , 0.05. b x2 32.707; degrees of freedom 4; P , 0.005.
The index of association (IA)1 has the form IA 5 100 3 c/(a 1 b 1 c), where c is the number of patches containing both channels, a is the number of patches containing only one type of channel, and b is the number of patches containing only the other type of channel. For the two fungal species we have examined, the IA is 58.0% (S. ferax) or 73.7% (N. crassa). The advantage of this technique is that it does not require knowledge of the number of patches lacking either channel or the actual number of channels in a patch. The disadvantage is that there is a possibility that the analysis can be misleading. This becomes clear if the two-way contingency test is used to analysis data from S. ferax. In a two-way contingency test, the observed numbers of patches containing both channel types, one or the other channel type, or no channels are compared to the expected values if the channels were distributed randomly. For S. ferax, a x2test of the observed and expected values reveals a highly significant association between the two channel types, but also a high proportion of K1 permeable channels which are always found alone, without stretch-activated channels (Table 4). The index of association is unable to reveal the presence of these two distinct K1 permeable channel populations. 1
Abbreviation used: IA, index of association.
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The relationship between the two channel types in the plasma membrane is clearly shown for S. ferax and N. crassa in Fig. 1. In S. ferax, there are two spatially distinct populations of K1 permeable channels. One population is found in the absence of stretch-activated channels, while the other population is always associated with stretchactivated channels. The K1 permeable/stretch-activated channel clusters are not disrupted by cytochalasin (Levina et al., 1994). By comparison, two spatially distinct K1 permeable channel populations are absent in N. crassa, and the association between stretch-activated channels and K1 permeable channels is weak, almost statistically nonsignficant. To summarize, analysis of distributions clearly shows that different types of ion channels in fungal plasma membrane exhibit distinct patterns of distribution: random, clustered, and/or associated with one another. At least in the case of S. ferax, nonrandom clustering of either channel type can be attributed to some actin-mediated connection to the cytoskeleton, although clusters of both channel types together must rely upon some other mechanism. The analysis up to now does not directly address asymmetric distributions of ion channels along the fungal hypha. Although x2tests can be used to analyze channel numbers in protoplasts from apical versus basal regions of the hypha (Garrill et al., 1992), the best determinant for asymmetric distribution relies upon direct examination of channel densities.
CHANNEL MAPPING Determining channel densities along a fungal hypha first requires knowledge of the surface area of the patch of
TABLE 4 Two-Way Contingency Test for Association of Spontaneous and Stretch-Activated Channels in Saprolegnia feraxa Spontaneous channels
Stretch-activated channels Present Absent
Present
Absent
40 (17.14) 24 (14.86)
5 (5.36) 15 (4.64)
Note. Data are from Levina et al. (1994). The observed and expected (in parentheses) numbers of observations are shown for presence and absence of channel type. a x2 59.26; degrees of freedom 2; P . 0.005.
Mapping Fungal Ion Channel Locations
73
membrane within the micropipette. The most accurate measurements can be performed using scanning electron microscopy, but unfortunately, this is a destructive assay. Instead, we have determined the relationship between bubble number (Mittman et al., 1987) and inside diameters measured from scanning electron micrographs and then used this linear regression to estimate patch area (Levina et al., 1994). The bubble number is a measure of the pressure required to pass air through the micropipette tip into methanol. Measurements of micropipette tip resistance may also be used, but the resistance varies with
FIG. 2. Channel densities along the hypha for Saprolegnia ferax. Stretch-activated channels are shown in the upper panel and spontaneous potassium permeable channels are shown in the lower panel for controls (circles) and cytochalasin E treatment (squares). The distance from the tip was calculated using the mean diameters of protoplasts originating from the hyphal tip, second protoplast from the tip, etc. The tip-high gradient of stretch-activated channels is statistically significant; the peak at the third protoplast from the tip for cytochalasin treatment is not. Data (mean 6 standard error, sample size ranged from 4 to 21) are redrawn from Levina et al. (1994).
FIG. 1. Relationship between stretch-activated (SA) calcium permeable and spontaneous potassium permeable ion channel densities for Saprolegnia ferax (top) and Neurospora crassa (bottom). Data are from Levina et al. (1994, 1995).
salt concentration in bathing and micropipette filling solutions, so errors are very likely. Channel densities can then be determined for apical to basal regions of the hypha. Results for S. ferax are shown in Fig. 2 and for N. crassa in Fig. 3.
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Roger R. Lew
ROLES IN TIP GROWTH
FIG. 3. Channel densities along the hypha for Neurospora crassa. Stretch-activated channels are shown in the upper panel and spontaneous potassium permeable channels are shown in the lower panel. Data (mean 6 standard error, sample size ranged from 3 to 10) are from Levina et al. (1995).
In the oomycete water mold S. ferax. there are tip-high gradients of spontaneous K1 permeable channels and stretch-activated Ca21 permeable channels (both gradients are statistically significant). Interestingly, actin disruption with cytochalasin E causes both tip-high gradients to disappear. This is strong evidence that the actin cytoskeleton controls channel location at the hyphal apex (Fig. 2). In the ascomycete N. crassa, there is no indication of a tip-high gradient of either the K1 permeable channels or stretch-activated channels (Fig. 3).
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Intuitively, if ion channels are explicitly required during the process of tip growth, one would expect them to be localized to the tip-growing region of the hypha. This is true in the case of the aquatic organism S. ferax. Potential roles for these channels in tip growth have been established (Garrill et al., 1993). Because the lanthinide gadolinium inhibits the stretch-activated channels, inhibits growth, and abolishes a tip-high intracellular Ca21 gradient observed during growth, the stretch-activated channels probably act as a growth sensor, That is, during tip expansion, activation of the channels would cause a localized influx of Ca21, which would act as a signal for continued secretion of membrane and cell wall material at the tip, thus maintaining tip expansion. The K1 permeable channels are inhibited by tetraethylammonium, which inhibits tip expansion transiently. Because the K1 movement is into the cell, the K1 permeable channels may function in maintaining intracellular osmolarity during the volume increase due to tip expansion. Because of the transient effect of channel inhibition, it is clear that the cell can utilize other mechanisms for maintaining intracellular osmolarity. In the terrestrial organism N. crassa, the absence of tip-high gradients of either channel type implies that the channels do not have a function in tip expansion. Indeed, the inhibition of the stretch-activated channels or K1 permeable channels with gadolinium or tetraethylammonium only transiently inhibits growth. It is possible that these striking differences between these two fungi are a result of evolution in two very different environments. In air, the fungal hyphal tip does not have an assured supply of extracellular Ca21 to rely upon as a component of a ‘‘growth-sensor’’ signal transduction system, and water required for volume increases may in fact have to travel considerable distances within the hypha to the growing tip. By contrast, evolution in an aquatic environment means that Ca21 and water are directly accessible to the growing tip. Hence, S. ferax would have no difficulty utilizing stretch-activated channels and K1 permeable channels in the tip growth process. Although there has been considerable research on channel location and clustering in animal neuronal cells (for a recent review, see Whatley and Harris, 1996), very few studies have been performed in nonanimal cells. Besides the work cited above using fungal organisms, Taylor et al. (1996) examined mechano-sensitive ion channel location in the fucus rhizoid. Similar to the results with N. crassa, they found no evidence for an asymmetric
Mapping Fungal Ion Channel Locations
localization of mechano-sensitive channels in these highly polarized algal zygotes (based on presence/absence criteria and channel density). In this system, Taylor et al. (1996) suggest that possible physiological functions for these channels could include osmoregulation and cytosolic Ca21 modulation of polar extension. Based upon the results to date, it is clear that there is a wealth of diversity in channel localization in the plasma membrane. Many questions regarding the extent to which this membrane architecture regulates cellular growth and morphological development remain to be explored. For nonanimal cells, the development of the laser ablation technique and its use in ion channel mapping in Fucus extend our knowledge to another kingdom besides the fungi. But within the fungal kingdom, there is a lot of work to be done.
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Roger R. Lew
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