From the * Department of Physiology and Department of Anesthesiology, University of California, Los Angeles, School of Medicine,
Los Angeles, California 90095
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ABSTRACT |
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When attached to specific sites near the S4 segment of the nonconducting (W434F) Shaker potassium channel, the fluorescent probe tetramethylrhodamine maleimide undergoes voltage-dependent changes in intensity that correlate with the movement of the voltage sensor (Mannuzzu, L.M., M.M. Moronne, and E.Y. Isacoff. 1996. Science. 271:213-216; Cha, A., and F. Bezanilla. 1997. Neuron. 19:1127-1140). The characteristics of this voltage-dependent fluorescence quenching are different in a conducting version of the channel with a different pore substitution (T449Y). Blocking the pore of the T449Y construct with either tetraethylammonium or agitoxin removes a fluorescence component that correlates with the voltage dependence but not the kinetics of ionic activation. This pore-mediated modulation of the fluorescence quenching near the S4 segment suggests that the fluorophore is affected by the state of the external pore. In addition, this modulation may reflect conformational changes associated with channel opening that are prevented by tetraethylammonium or agitoxin. Studies of pH titration, collisional quenchers, and anisotropy indicate that fluorophores attached to residues near the S4 segment are constrained by a nearby region of protein. The mechanism of fluorescence quenching near the S4 segment does not involve either reorientation of the fluorophore or a voltage-dependent excitation shift and is different from the quenching mechanism observed at a site near the S2 segment. Taken together, these results suggest that the extracellular portion of the S4 segment resides in an aqueous protein vestibule and is influenced by the state of the external pore.
Key words: Shaker potassium channel; fluorescence quenching; pore conformational changes ![]() |
INTRODUCTION |
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The Shaker potassium channel, which has served as a
model for understanding the behavior of voltage-gated
ion channels, opens in response to depolarized potentials to allow potassium flow (Timpe et al., 1988).
Opening and closing of the channel is controlled by
the channel's voltage sensor, whose movement is reflected by the gating current (Armstrong and Bezanilla,
1973
; Bezanilla et al., 1991
). Recent studies have shown
that this gating current is generated in part by the
movement of charged residues in the S2 and S4 segments across the transmembrane electric field (Seoh et al., 1996
; Aggarwal and MacKinnon, 1996
). Conformational changes associated with the voltage sensor
have also been measured using extrinsic fluorescent
probes attached to specific sites near the S4 segment
(Mannuzzu et al., 1996
; Cha and Bezanilla, 1997
).
Previous fluorescence studies of proteins have used
measurements of anisotropy and quencher accessibility
to answer questions about the fluorophore's environment (reviewed by Eftink, 1991). Fluorescence anisotropy measures the rotational mobility of the fluorophore and reflects the fluorophore's environmental
constraints. For instance, the anisotropy of fluorophore
dissolved in glycerol is very high, consistent with a viscous environment. Fluorescence-quenching studies with
various molecules such as D2O and iodide have been used to examine fluorophore exposure to the aqueous
environment. By modulating the state of the protein
and measuring changes in fluorescence quenching, state-specific accessibilities of particular residues can be determined.
Understanding the mechanism of fluorescence quenching can also yield information about the fluorophore's
environment. For instance, changes in fluorescence intensity can be caused by reorientation of the fluorophore's transition dipole, leading to a concomitant change in absorption cross-section (Andreev et al.,
1993). Shifts in the excitation spectrum of the dye,
which can be caused by fluorophore-fluorophore interactions, can lead to changes in absorption at particular excitation wavelengths and a corresponding change in emission (Burghardt et al., 1996
). Changes in the hydrophobicity of the fluorophore's environment can
lead to changes in fluorophore quenching. Finally,
nearby protein residues can also interact with and
quench the fluorophore. By delineating the specific
mechanism of quenching in the Shaker potassium channel, a better understanding of the conformational
changes near the S4 segment can be obtained.
We report here that the fluorescence changes in the extracellular region of the S4 segment are affected by different substitutions in the pore. This result was unexpected because interactions between the S4 segment and external pore have not previously been observed. The external application of tetraethylammonium (TEA)1 and agitoxin to a conducting version of the channel also affects the fluorescence quenching through an interaction with the pore. This effect is seen as the elimination of a fluorescence component whose voltage dependence coincides with ionic activation, but whose kinetics are slower. This result suggests that there are conformational changes coupled to channel opening that affect the extracellular portion of the S4 segment and are blocked by TEA or agitoxin.
To better interpret these results, we turned to other
techniques to determine the environmental properties
of the fluorophore. The modulation of the fluorescence by pH and collisional quenchers, along with
anisotropy measurements, indicates that the fluorophore may be interacting with a pH-titratable protein
vestibule. This idea is supported by a recent study that
suggested that narrow vestibules that line the S4 segment permit the passage of protons but exclude cysteine-reactive reagents (Starace et al., 1997). These experiments necessitated the development of a new optical technique based on an upright microscope and a
water-immersion objective. This technique enabled measurements of fluorescence polarization and increased
the efficiency of light collection by a factor of >10 over
the previous cut-open oocyte epifluorescence setup.
Although the idea of fluorescence quenching by protein residues was substantiated by these results, several other mechanisms that do not involve protein quenching were tested as possible mechanisms for the signal seen near the S4 segment. Although the quenching mechanism near the S4 segment does not appear to involve a reorientation of the fluorophore, a change in environmental hydrophobicity, or a voltage-dependent excitation shift, a site near the S2 segment does undergo a voltage-dependent excitation shift. Thus, other regions of the protein may undergo different changes in environment.
By combining information from the study of the W434F and T449Y Shaker constructs with other measurements, we propose that the voltage-dependent fluorescence quenching of tetramethylrhodamine maleimide (TMRM) near the S4 segment is modulated by the state of the external pore. In addition, properties of the fluorophore indicate the presence of a nearby protein vestibule that lines the extracellular region of the S4 segment.
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MATERIALS AND METHODS |
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Modified Cut-Open Oocyte Epifluorescence Setup
The measurement of gating currents were made using the cut-open oocyte technique for spatial voltage homogeneity and fast temporal resolution (Stefani et al., 1994). To make polarization and anisotropy measurements possible and improve the efficiency of light collection, the experimental setup was modified
from an inverted microscope with fiber optic (Cha and Bezanilla,
1997
) to an upright microscope with a water-immersion objective
(Fig. 1). Using this microscope, the intensity of light, as measured at the photodiode or at the CCD, achieved levels more
than 10× what was measured with the fiber optic configuration.
This difference was due in part to light loss from the coupling between the fiber optic and the oocyte surface, and also from the
coupling between the inverted microscope objective and the fiber optic. The water-immersion objective's field of view matched
the 600-µm-diameter oocyte surface visible in the top cut-open
chamber, and the objective's numerical aperture was larger than
that of the optics used in the inverted microscope setup.
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The upright microscope's narrow base permitted the placement of the cut-open oocyte chamber directly beneath the objective. The chamber was placed on a set of sliders so that the oocyte could be mounted and permeabilized under a dissection microscope, and then moved directly underneath a 4× objective for insertion of the microelectrode, and then a 40× objective for optical measurements.
The optical setup consisted of a BX50WI microscope (Olympus Optical, Melville, NY) and used excitation filters, dichroic mirrors, and emission filters (Omega Optical, Brattleboro, VT, and Chroma Technologies, Brattleboro, VT) appropriate for tetramethylrhodamine-5-maleimide (Molecular Probes, Eugene, OR). A microscope tungsten lamp (Carl Zeiss Corp., Thornwood, NY) with a 150-watt filament, powered by a 6286A power supply (Hewlett Packard, Palo Alto, CA), served as the light source. The lamp output was interrupted with a TTL-triggered VS25 shutter (Vincent Associates, Rochester, NY) to minimize photobleaching of the probe.
The LUMPlanFl 40× water-immersion objective had a numerical aperture of 0.8 and working distance of 3.3 mm (Olympus
Optical). Light measurements were made with a PIN-020A photodiode (UDT Technologies, Torrance, CA) mounted on an FP-1
fiber optic manipulator (Newport Corp., Irvine, CA), which was
attached to the front end of an optical splitter at the microscope's epifluorescence port. The photodiode was attached to
the headstage input of an integrating patch clamp amplifier for
low noise amplification of the photocurrent. The patch clamp
amplifier was an Axopatch-1B (Axon Instruments, Foster City,
CA), used with an IHS-1 integrating headstage. A circuit with a
45-volt battery (Eveready, St. Louis, MO) and a 10-G resistor
was used to remove integration spikes by offsetting current into
the summing junction of the headstage. The fluorescence emission was focused onto the photodiode active area using a microscope condenser lens with a focal distance of 1 cm.
The voltage clamp setup was composed of a top, middle, and bottom chamber (Fig. 1, bottom). The bottom chamber contained the portion of the oocyte that was permeabilized with saponin so that current could be injected directly into the oocyte. The middle chamber served as an electronic guard, and the top chamber, which was painted black, contained the portion of the oocyte membrane from which the fluorescence changes and gating or ionic currents were measured. The voltage electrode measured the membrane potential across the oocyte membrane and was part of the feedback loop that held the interior of the oocyte at virtual ground. Voltage clamp of the oocyte was performed with a CA-1 cut-open oocyte clamp (Dagan Corp., Minneapolis, MN).
Anisotropy and Polarization Measurements
The use of an upright microscope and water-immersion objective
enabled measurements of polarization and anisotropy that were previously impossible because the fiber optic used to image the oocyte did not maintain light polarization. With rotatable polarizers in the excitation and emission pathway (Olympus U-AN360;
Olympus Optical), two measurements of anisotropy were possible for each polarizer, one in a north-south orientation with respect to the microscope, and the other in an east-west orientation. We measured the fluorescence of labeled, expressing oocytes
using all four possible combinations of excitation and emission
polarizers: exciter north-south polarized, and emitter either north-
south (parallel, I) or east-west (perpendicular, I
) polarized; or
exciter east-west polarized, and emitter either east-west (parallel, I
) or north-south (perpendicular, I
) polarized. By measuring the intensity of fluorescence polarized parallel (I
) and perpendicular (I
) to the excitation light, the steady state anisotropy
A of the fluorophore can be calculated using the equation A = (I
I
)/(I
+ 2I
), where A ranges between 0 in the completely isotropic case and 0.4 in the completely anisotropic case
(Cantor and Schimmel, 1980
). Using the four polarization measurements, two independent calculations of anisotropy can be made. With correction factors to account for the intrinsic polarization properties of the optical path, both calculations should
yield the same value of anisotropy, independent of excitation polarization.
The calibration process was done by measuring the anisotropy of a known system, TMRM dissolved in glycerol, and then calculating the correction factor for the north-south and east-west excitation polarizations, which would give the correct anisotropy value from the actual microscope measurements. The anisotropy of TMRM in glycerol is 0.38 (P. Selvin, personal communication); the correction factor was 0.971 in the north-south excitation polarization and 1.312 in the east-west excitation polarization.
The contribution of autofluorescence and fluorescence not arising from channels was quantified by measuring the mean fluorescence intensity from labeled populations of channel-expressing oocytes and comparing them to the fluorescence intensity of labeled and unlabeled nonexpressing oocytes. The results are shown in Table I.
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Because the average fluorescence intensity of a labeled, expressing oocyte is typically 6-10× that of a labeled, uninjected oocyte, >85% of the fluorescence is coming from labeled channels. With this data, we have assumed that the anisotropy, quenching, and polarization measurements primarily reflect the properties of fluorophores attached to the channel.
Spectral Analysis
Spectra were obtained with a Multispec 257i spectrograph (Oriel
Instruments, Stratford, CT) with a 1,200 lines/mm grating for
100 nm bandwidth, attached to an Instaspec V CCD camera with image intensifier cooled to 20°C (Oriel Instruments). The spectrograph and CCD were attached to the rear end of an optical
splitter at the epifluorescence port of the microscope.
The filter response of the dichroic mirror was corrected by obtaining the transfer function of the mirror. This was calculated by taking the spectrum obtained by transillumination of the tungsten lamp with mirror and dividing it by the spectrum obtained by transillumination of the lamp alone. Spectra of TMRM in different solvents were measured by dissolving TMRM to a 5-µM concentration, and the measurements were taken with a 535DF35 excitation filter and a 570DRLP dichroic mirror (Omega Optical). Because of the shallow cutoff of the dichroic mirror, spectral characteristics of the signal were maintained to <560 nm with this procedure.
Data Acquisition and Analysis
Gating, ionic, and fluorescence currents were acquired with a PC44 board (Innovative Technologies, Moorpark, CA), which interfaces with a Pentium-based computer via an IBM-compatible AT slot. The fluorescence and electrophysiology were simultaneously acquired on two 16-bit analogue-to-digital converters and transferred to two separate channels of the PC44. When data is sampled at intervals longer than 5 µs (all traces presented in this paper), the program running the PC44 board acquires the data at 5 µs per point, and then decimates the data to the required sampling period after digitally filtering the original data to the new Nyquist frequency. The acquisition program and data analysis programs were developed in house and were run in MS-DOS and Windows 95, respectively.
Molecular Biology, Channel Expression, and Oocyte Labeling
The noninactivating (6-46, IR) version of the Shaker H4 channel (H4IR) was originally cloned into an engineered version of the
pBSTA vector. Two different vector backgrounds were used: a
nonconducting version of the channel (W434F), and a conducting version of the channel that tightly binds TEA and agitoxin 2 (T449Y). The agitoxin 2 was kindly provided by Dr. Adrian Gross
(UCLA, Los Angeles, CA). For site-directed mutagenesis of all
constructs, a two-step PCR protocol (Moore, 1994
) was used to introduce mutations between the XbaI and BglII sites into the
Shaker background. After subsequent cloning into the pBSTA vector, the cDNA generated by PCR was sequenced to exclude the
possibility of unwanted mutations. The constructs are designated
by the original amino acid, residue number, and substituted amino
acid (i.e., M356C designates the construct where cysteine was substituted for methionine at residue 356).
The cRNA was transcribed in vitro with the T7 mMessage machine kit (Ambion Inc., Austin, TX), and 50 nl cRNA at a concentration of 100 ng/µl were injected into each Xenopus oocyte. Experiments were performed from 2 to 7 d after injection. The sterile oocyte incubation solution consisted of 100 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 1 mM MgCl2, 5 mM HEPES, 10 µM EDTA, and 100 µM dithiothreitol.
For fluorescent labeling, Xenopus oocytes were incubated in a
depolarizing solution containing 5 µM tetramethylrhodamine-5-maleimide (Mannuzzu et al., 1996) at 18°C for 40 min.
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RESULTS |
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Modulation of Fluorescence Quenching near the S4 Segment by Substitutions in the External Pore
By substituting cysteine for a specific residue, site-directed
fluorescent labeling can be used to measure site-specific environmental changes with the covalent attachment of an extrinsic, membrane-impermeant fluorescent probe to the introduced cysteine (Mannuzzu et
al., 1996; Cha and Bezanilla, 1997
). Using this technique, sites M356C and A359C, located in the extracellular portion of the S4 segment (as a reference, R362 is
the outermost charged residue in the S4 segment), displayed voltage-dependent fluorescence changes with
kinetic and steady state properties similar to those of
the gating currents. Thus, the fluorescence changes near
the S4 segment appear to reflect conformational changes
associated with movement of the voltage sensor.
The properties of the fluorescence signal measured at sites near the S4 segment are also affected by mutations in the external pore. This result was surprising because it has not been shown that the S4 segment is affected by the state of the external pore. In the fluorescence traces for the M356C construct combined with the T449Y mutation in the external pore (M356C T449Y), there is a slow fluorescence component that is visible at large depolarizations and is absent in the M356C construct combined with the W434F mutation in the external pore (M356C W434F) (Fig. 2 A). This component is likely responsible for the shallow voltage dependence seen in the F-V curve of the M356C T449Y construct, in contrast to the M356C W434F construct (Fig. 2 B).
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Fluorescence signals from site A359C are also affected by pore substitutions: the kinetics for the A359C T449Y construct are considerably different than for the A359C W434F construct (Fig. 2 C). In this case, the fluorescence kinetics appear to be faster for A359C T449Y than for A359C W434F. The F-V curve also reflects a shallow voltage dependence of the fluorescence changes in the A359C T449Y construct in comparison with the A359C W434F construct (Fig. 2 D).
The mutations W434F and T449Y, both located in
the external mouth of the pore, have very different effects on conduction. The tryptophan-to-phenylalanine
mutation at residue 434 (W434F) prevents conduction
without affecting the conformational changes that occur in the internal mouth of the pore, as judged from the effects of internal TEA on the gating currents (Perozo
et al., 1993). In comparison, the threonine-to-tyrosine
mutation at residue 449 (T449Y) preserves ionic conduction and increases the affinity of ionic blockers such
as TEA and agitoxin (MacKinnon and Yellen, 1990
;
Heginbotham and MacKinnon, 1992
; Gross and MacKinnon, 1996
). The differences between these two constructs could be attributed either to changes in the
movement of the voltage sensor or changes in the fluorophore's environment introduced by the pore mutations. A comparison of the gating currents of conducting and nonconducting (W434F) constructs indicate
that the voltage sensor properties do not appear to be
altered by this pore substitution (Perozo et al., 1993
).
Thus, one possible explanation for the effect is that
these pore mutations modify the channel structure so
that a different optical profile is seen by the fluorophore. Another possibility is that the state of the external pore may be coupled to the state of the S4 segment
in a manner that is not easily observed in the gating
currents. A third explanation, which will be addressed
in the next section, is that the presence of ion flow
through the channel directly affects the fluorophore.
Effects of TEA and Agitoxin on Fluorescence Quenching
To determine whether ion flow through the channel affects fluorophore properties near the S4 segment, we compared the fluorescence signals in the M356C T449Y construct before (Fig. 3 A) and after (Fig. 3 B) the application of external 120 mM TEA-Mes to block the outward flow of potassium. The fluorescence traces are superimposable at small depolarizations but diverge for larger depolarizations. This is more clearly seen in a plot of the fluorescence change versus voltage, or F-V curve (Fig. 3 C). A similar effect in the M356C T449Y construct was seen when ionic current was blocked by the addition of 3 µM agitoxin to the external solution (Fig. 3, D-F). In both cases, the fluorescence quenching became much larger after blocking ionic flow.
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Although the characteristics of the fluorescence signal are somewhat different, a similar effect of TEA or agitoxin is seen at a second site, A359C, which lies just three residues closer to the S4 segment (Fig. 4). Again, the fluorescence signals are superimposable at small depolarizations, but become larger at potentials where the channel has opened (Fig. 4, C and F). These effects do not appear to be related to series resistance error, as the effect is consistently observed in oocytes independent of the size of the ionic currents.
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The effects of TEA or agitoxin on the fluorescence at both sites can be obtained by subtracting the fluorescence signals before and after blocker application to obtain the fluorescence difference (dF, see Fig. 5). This effect, as measured by the fluorescence difference before and after the addition of pore blocker, displays a voltage dependence (dF-V) that coincides with ionic conductance, or channel activation (G -V).
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It is unlikely that this modulation is due to a direct interaction of TEA or agitoxin and the fluorophore. Adding either TEA or agitoxin to either W434F construct (M356C, A359C) does not affect the fluorescence signal (data not shown). However, it is possible that the fluorophore is being directly affected by potassium flux. If this were the case, we would expect the following four results: first, the voltage dependence of potassium-based quenching should follow the current-voltage, or I-V curve, and consequently should remain linear at depolarizing potentials. Second, the application of TEA or agitoxin should decrease the quenching of the fluorophore at depolarized potentials by reducing outward potassium flow. Third, the time course of the fluorescence difference should follow the kinetics of ionic current. Fourth, when the direction of ionic current reverses, the fluorescence intensity should also reverse and show substantial intensity changes that mirror the direction of ionic flow.
We found that none of these results apply to the measured fluorescence difference. First, the voltage dependence of the fluorescence difference saturates at large
depolarizations (Fig. 5). Second, the application of
TEA or agitoxin actually increases the quenching seen
at depolarized potentials (Figs. 3 and 4). To address
the third point, we compared the kinetics of the fluorescence difference to ionic current difference. Although the voltage dependence of this signal matches
the voltage dependence of ionic conductance (Fig. 5),
the kinetics of this signal are significantly slower than
those of ionic activation (Fig. 6). For three different potentials for each combination of site and blocker, the
ionic current difference and fluorescence difference
for each potential were superimposed by normalizing
to the final value of a 40-ms pulse from 90 mV. From
this comparison, it is apparent that the fluorescence
signal is generally slower than the ionic current difference at these potentials for each site and blocker. With
regard to the fourth point, when the ionic current reverses direction during repolarization (Figs. 3 and 4, insets), the fluorescence does not demonstrate a similar
reversal or increase from the initial intensity level. Instead, the fluorescence decays back to its original level,
suggesting that the fluorescence is not directly affected
by the direction of ionic flow. Taken together, these observations argue that TEA and agitoxin do not modulate fluorescence by the presence or absence of ion
flow. Instead, TEA and agitoxin seem to modulate fluorescence by a mechanism related to their ability to prevent conduction. These molecules may inhibit conformational changes that normally occur in the outside region of the conducting channel. These conformational changes, which are coupled to channel opening, may
modulate the fluorescence quenching seen at sites near
the S4 segment.
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Modulation of Fluorescence by pH Indicates Possible Interactions with pH-titratable Residues
One of the implications of the previous experiments is that the fluorophore may be interacting with another region of the protein. To determine whether the fluorophore may be quenched by nearby protein residues, we examined the characteristics of the fluorophore environment by modulating the external pH. Because TMRM's quantum yield does not change with pH, modulation of TMRM fluorescence by pH would indicate interaction with the probe with a nearby protein whose properties change with pH. The lack of effect of pH on TMRM alone was confirmed by determining that the background fluorescence of uninjected, labeled oocytes does not change with external pH (data not shown). Thus, changing the pH should not directly modulate the fluorescence signal from a labeled site on the protein.
Nevertheless, changing the pH shifts the gating
charge versus voltage, or Q-V curve, due to changes in
the surface charge detected by the voltage sensor (Starace et al., 1997). Because fluorescence changes at sites
near the S4 segment reflect properties of the voltage
sensor, the F-V curves at sites M356C and A359C as a
function of pH should reflect this shift caused by surface charge. Fig. 7 illustrates the changes that occur in
the fluorescence intensity versus voltage curves as a
function of pH. As expected, the curve is shifted by the
external pH, with pH 5.1 corresponding to the curve
shifted most to depolarized potentials.
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However, in addition to the shift, there is a difference in the magnitude of the TMRM intensity as a function of pH at both sites, indicating that pH does modulate the voltage-dependent quenching. The increase in fluorescence intensity is largest at hyperpolarized potentials and acidic pH values. Because TMRM's quantum yield is not directly affected by pH, it appears that pH affects the properties of the quencher that interacts with the probe. In addition, both sites show a similar increase in intensity at more acidic pH values. This indicates that the fluorophore at either site may be interacting with a similar region of protein.
This leads to two possible hypotheses: pH may affect the global conformation of the channel so that the probe interacts with a different region of the channel, or pH may affect the nearby region of protein, which interacts directly with the probe. The global effect of pH, as seen by a shift in the voltage axis, results from the change in surface charge. If this change in surface charge were responsible for conformational changes that modulate the fluorescence, then other methods that change the surface charge should have a similar effect. Increasing the concentration of external calcium mimics the surface charge effects of acidic pH and shifts the Q-V curve to more depolarized potentials. However, changing the calcium concentration from 1.8 to 20 mM shifts the F-V curve without modulating the magnitude of the fluorescence at any potential (Fig. 8). This implies that the fluorescence quenching of the probe is independent of changes in surface charge, which supports the idea that there is a region of protein that interacts directly with the fluorophore. This pH effect could either be direct (nearby residues quench differently because of a change in charge) or indirect (the fluorophore moves to a different environment because of electrostatic changes caused by pH).
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Differential Access of Collisional Quenchers Indicate the Presence of an Aqueous Protein Vestibule Surrounding the Extracellular Portion of the S4 Segment
If the fluorophore were interacting with a nearby region of protein that acts as a quencher as a function of
voltage, one might expect that accessibility to the fluorophore would also be constrained by this region of
protein as a function of voltage. Thus, using quenchers
with different properties to examine residue accessibility could yield information about nearby structural rearrangements (Arias, 1993; Eftink, 1994). Iodide, a
negatively charged anion, quenches sites that are exposed to solvent in an electrostatically dependent manner. In comparison, D2O is a much smaller quencher
whose substitution for H2O typically increases fluorophore intensity at solvent-exposed sites. We measured
the effects of these solvents on free TMRM and determined that its intensity decreases in the presence of iodide and increases in the presence of D2O (Fig. 9).
With this knowledge, we looked at the accessibility of
specific residues to both quenchers as a function of
voltage.
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We first examined the accessibility of TMRM at positions M356C and A359C to external iodide. The change in fluorescence caused by quencher was measured as the intensity ratio (R) of fluorescence intensity measured after addition of quencher to the fluorescence intensity measured before addition of quencher (Fig. 10). If the fluorescence intensity after quencher application is less than the fluorescence intensity before quencher application, then R < 1. At both sites near the S4 segment, R < 1 (P < 0.01), indicating that iodide is effective in quenching the TMRM fluorescence at both sites and therefore has access to these sites (Fig. 10 A, left).
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To determine the relative accessibility to iodide as a
function of potential, the ratio of R at 90 mV (R
90 mV)
to R at 0 mV (R0 mV) was calculated (Fig. 10 A, right). Because the voltage was held at 0 mV, the channels are
likely in the slow-inactivated state. At site M356C, R
90 mV:
R0 mV > 1 (P < 0.05), indicating that iodide appears to
have better access at depolarized potentials.
The accessibility to D2O at both sites was determined
using the same ratios (Fig. 10 B). Since R > 1, the fluorescence intensity increases after the application of
D2O at both sites (P < 0.01) (Fig. 10 B, left). Thus, both
sites also appear to be accessible to water at 90 and
0 mV. This makes it highly unlikely that the fluorescence change is caused by movement of the fluorophore from a completely hydrophobic environment
into the aqueous environment. This was also previously
inferred from spectral analysis (Cha and Bezanilla, 1997
).
The ratio of R at 90 mV to R at 0 mV was also calculated for D2O. At site A359C, R
90 mV:R0 mV < 1 (P < 0.01), indicating better access at depolarized potentials. However, M356C shows a value >1 (P < 0.01), indicating that D2O access is favored at hyperpolarized
potentials. At first, this result appears to contradict the
result obtained with iodide. But the differential access
can be explained by the properties of these quenchers.
Because iodide is negatively charged, it can show preferential access dependent on electrostatic properties,
in contrast to water. Thus, M356C may see a more negatively charged environment at hyperpolarized potentials, which would explain the decreased iodide access,
while seeing a larger crevice, which would explain the
greater D2O accessibility. Similarly, M356C could also
see a more positively charged, but smaller crevice at depolarized potentials.
Differences in Anisotropy Are Consistent with Protein Constraints Near the S4 Segment
Fluorescence anisotropy is a reflection of the rotational
freedom of the fluorophore during its excited-state lifetime. If the fluorophore were surrounded by protein,
one might also expect that anisotropy of the fluorophore would be affected by constraints imposed by
nearby residues. To examine possible constraints on the mobility of the fluorophore, the steady state anisotropy values of TMRM attached to different sites in the
channel were measured as a function of holding potential (Fig. 11 A). The measured anisotropy is lowest at
the site near the S2 segment (D270C), whereas, near the S4 segment, the anisotropy is lowest at residue
V363C and is larger at sites M356C and A359C, regardless of pore mutation. One view of the S4 segment is
that some residues in the S4 segment are relatively buried, while more extracellular residues in the S4 segment appear to be more accessible (Larsson et al.,
1996). This data is the opposite of what would be expected if residues in the S4 segment were constrained
by protein or lipid, and residues outside the S4 segment lie further from these constraints. In addition,
this profile implies that this region near the S4 segment may lie in close proximity to another region of protein
that affects the anisotropy of TMRM.
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These anisotropy values change slightly in response
to voltage, and the direction of the change is dependent on the site. The ratio was taken of the anisotropy
at 90 mV (A
90 mV) to the anisotropy at 0 mV (A0 mV)
(Fig. 11 B). At site M356C, the fluorophore shows
larger anisotropy at depolarized potentials (P < 0.01),
whereas at sites A359C (P < 0.01) and V363C (P < 0.01), the anisotropy decreases at depolarized potentials. This implies that the environment near residue
M356C becomes more constrained at depolarized potentials, which is consistent with the results from D2O
exposure. Taken together, these results are not consistent with a model where the S4 segment simply moves
from within the lipid bilayer into the aqueous environment as the channel opens. It is more likely that all
three sites lie in a vestibule surrounded by protein, and
some residues may experience relief from nearby protein
constraints at depolarized potentials (A359C, V363C), while others may not (M356C).
Changes in Fluorophore Orientation Are Not Responsible for the Fluorescence Quenching
To test the hypothesis that the voltage-dependent fluorescence changes are due to quenching by a nearby region of protein, we explored two other mechanisms that could explain the fluorescence changes. The first mechanism we tested is a voltage-dependent reorientation of the fluorophore. With polarized excitation light, only those fluorophores with transition dipoles oriented parallel to the polarization of incoming light will be excited; with a polarized emission filter, only the fluorophores that emit light with that polarization will be seen. The change in fluorescence intensity could be caused by a reorientation of the dipole: if the incoming light has polarization properties, which is typical for an epifluorescence setup with a dichroic mirror, then a reorientation of the fluorophore's dipole could change the relative absorption of one incoming polarization with respect to another. This polarization shift could modulate the total intensity of the fluorophore emission as well as the relative polarization intensities of emitted fluorescence.
If the change in fluorescence intensity is caused by a change in orientation of the fluorophore and corresponding transition dipole, then the decrease in emitted fluorescence at one excitation polarization should be accompanied by an increase in fluorescence at the perpendicular polarization. To determine whether a change in polarization was responsible for the voltage-dependent fluorescence changes, the F-V curves at sites M356C and A359C were measured at the four possible orientations of excitation and emission polarization. When examining the normalized change in fluorescence as a function of polarization at these sites, the direction and voltage dependence of fluorescence change is maintained at all possible polarizer orientations, indicating that the fluorescence quenching is not caused by changes in orientation of the fluorophore (Fig. 12). Because the fluorescence signal does not change directions as a function of polarization, it cannot be primarily responsible for the voltage-dependent fluorescence change.
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A Voltage-dependent Excitation Shift Is Responsible for Fluorescence Quenching Near the S2 Segment, but Not Near the S4 Segment
Another possible source of quenching could be the result of a shift in the absorption spectrum, which can be
caused by the interaction of neighboring fluorophores
and does not typically involve protein quenching (Selwyn and Steinfeld, 1972; Burghardt et al., 1996
). To examine the possibility of a shift in the excitation of the
fluorophore as a function of voltage, we examined
changes in fluorescence as a function of the excitation
wavelength. In principle, with a shift of the excitation
spectrum, the direction of fluorescence change when
exciting on one side of the excitation peak should be
the opposite of the direction of the fluorescence change when exciting on the other side of the peak.
Fig. 13 A presents simulated excitation and emission
spectra to illustrate how this shift in excitation can induce changes in emission. For instance, if the excitation peak shifts to shorter wavelengths when the membrane is depolarized from
90 to 0 mV, then at excitation wavelengths
1, the fluorescence will increase,
whereas, at excitation wavelengths
2, the fluorescence will decrease.
|
By using different filters to excite several regions of
the excitation spectrum of TMRM, we can determine
whether the fluorescence change is caused by an excitation shift by looking for a direction reversal of the fluorescence change as a function of excitation wavelength.
At site D270C, which is located in the extracellular portion of the S2 segment, there is a small decrease in fluorescence (~0.5% F/F), denoted by an upward deflection of the trace, in response to a depolarizing pulse to
0 mV when illuminating with light between 510 and
560 nm (Fig. 13 B). In comparison, when illuminating with light between 453 and 487 nm, the fluorescence
increases for the same depolarization. This indicates
that different wavelengths excite fluorescence on different sides of an excitation peak that shifts in response
to voltage. Fluorescence changes were not previously observed at this site because of insufficient optical sensitivity of the setup.
We then examined the normalized fluorescence change as a function of voltage, or F-V curve, at sites M356C and A359C centered at three different wavelengths: 557, 535, and 450 nm. For all filters, the fluorescence decreases in response to depolarizations, and the F-V curve is superimposable at all three excitation wavelengths. This result indicates that there is no excitation shift at either site (Fig. 14). Thus, different fluorescence quenching mechanisms occur in different regions of the channel, and the small fluorescence changes at site D270C arise in a different manner than the large fluorescence changes near the S4 segment. Because the voltage-dependent fluorescence changes in the extracellular region of the S4 segment are not caused by an excitation shift of TMRM, it is unlikely that the underlying mechanism is related to fluorophore-fluorophore interactions, such as dimer formation.
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DISCUSSION |
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Implications for Conformational Changes in the External Pore
The fluorescence signal near the S4 segment, which was previously thought to be an indicator of conformational changes specific to the S4 segment itself, is also modulated by mutations and blockers of the external pore. In particular, the conducting T449Y construct shows fluorescence changes with very different characteristics than the nonconducting W434F construct, and the T449Y fluorescence signal changes dramatically with the application of the pore-blocking molecules TEA or agitoxin (Fig. 3). Because this effect is not seen in the W434F construct, these molecules must modulate fluorescence by affecting the state of the pore. This modulation could occur because inhibition of conductance in the external pore could propagate back through the activation pathway and affect the movement of the S4 segment. The other possibility is that the fluorophore's environment contains residues that are affected by the external state of the pore, with the implication that the S4 segment may lie in close proximity to the external pore.
The effect of these blockers, as measured by the fluorescence difference before and after application of
blocker, appears to share the voltage dependence of
ionic activation, albeit with slower kinetics (Fig. 6).
These slower kinetics indicate that the conformational
changes represented by these traces are not responsible for directly gating ion flow. But the changes are likely coupled to the open state of the channel and may
serve as preparatory steps toward slow inactivation. Similar results were obtained in experiments that identified fluorescence changes at sites F425C and T449C in
the external pore, which were slower than ionic activation (Cha and Bezanilla, 1997).
The fluorescence difference also has implications for the structure of the W434F nonconducting construct, whose fluorescence signal is unaffected by TEA or agitoxin. The absence of modulation indicates that the W434F construct may be unable to undergo these external conformational changes near the pore, or that the W434F construct may not effectively bind these blockers. In fact, both hypotheses could hold true, and the mutation may change the pore structure in a manner that prevents conformational changes and reduces TEA and agitoxin affinity. In either case, the W434F mutation alters the structure of the external pore such that the characteristics of the quenching near the S4 segment are different than those seen in the T449Y-conducting construct.
Mechanisms of Fluorescence Quenching in the Shaker Potassium Channel
By studying the mechanism of fluorescence quenching
near the S4 segment, we hoped to elucidate the characteristics of the environment surrounding these residues. To this end, several potential quenching mechanisms were ruled out as being responsible for the large
voltage-dependent changes in fluorescence intensity.
Polarization and excitation shift experiments indicate
that neither a reorientation of the fluorophore nor a
shift of excitation wavelengths are responsible for the
fluorescence change. In addition, access to the fluorophore by D2O at 90 and 0 mV argues strongly against
a movement of the fluorophore from a completely hydrophobic environment into an aqueous environment.
Thus, the remaining quenching mechanism, voltage-dependent quenching by nearby protein residues, becomes the most likely candidate.
Several other observations support protein-based
quenching as the actual mechanism. Modulation of
fluorescence by the state of the external pore suggests
that the fluorophore may interact with residues affected by the state of the pore. In addition, anisotropy
measurements indicate that the environment near the fluorophore becomes more constrained at sites near
the S4 segment that show a large fluorescence change
(M356C, A359C) than at sites near the S2 segment
(D270C) or in the S4 segment itself (V363C), which
show little or no fluorescence change. Experiments examining pH titration and the relative accessibility of
different quenchers suggest the existence of nearby
electrostatic and steric constraints, which are best explained by interactions with a nearby protein vestibule.
Thus, the voltage-dependent quenching near the S4
segment appears to depend on quenching by nearby
protein residues, which has been seen in other systems
(Conibear et al., 1996; Coelho-Sampaio and Voss, 1993
).
Properties of a Putative Protein Vestibule Near the S4 Segment
An extracellular protein vestibule near the S4 segment
has been proposed, based on histidine scanning mutagenesis (Starace et al., 1997). The experiments presented in this paper can be used to visualize characteristics of residues lining part of this vestibule. An important caveat is that the properties of the environment of
residues near the S4 segment have been inferred from
characteristics of the fluorophore, which is attached to
the residue of interest with a maleimide linker. With a
typical length of ~7 Å, the cysteine-reactive maleimide
linker acts as a tether that cannot completely constrain the position of the fluorophore near the residue. The
implications of structural features near the labeled site
must be made with the knowledge that these structures
lie within the reach of a molecule that contains both
fluorophore and linker (see Table II).
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With this in mind, general characteristics of the protein environment can be inferred from the anisotropy, pH, and collisional quencher studies. For instance, given the higher anisotropy values at sites M356C and A359C than at site V363C, it appears that the vestibule may be narrower further into the S3-S4 linker and wider near the beginning of the S4 segment. In addition, anisotropy studies indicate that M356C becomes more constrained at depolarized potentials, indicating that the vestibule is still sensed at depolarized potentials. D2O accessibility studies indicate that the vestibule is aqueous at all sites and a broad range of potentials. The increased accessibility to iodide at depolarized potentials at site M356C indicates that the environment at this site may be positively charged at depolarized potentials, or negatively charged at hyperpolarized potentials.
If the probe is quenched by a residue or group of residues whose ability to quench is pH dependent, likely
candidates include aspartic and glutamic acid (pKa
4-5), as judged from the marked effects and lack of saturation near pH 5 (Fig. 7). Thus, the pH titration data,
which indicates possible interaction with glutamic or
aspartic acid at hyperpolarized potentials, are consistent with the hypothesis that the fluorophore may lie in
a negatively charged environment in the closed state of
the channel.
This information, summarized in Table II, can be
used to create a preliminary picture of the vestibule
near the S4 segment (Fig. 15). This illustration highlights the distinctive properties of these three sites at
90 and 0 mV, particularly at site M356C, which behaves differently than the other sites. This diagram
models the movement of the S4 segment as a change in
orientation, or tilt, with respect to the transmembrane
field (Papazian and Bezanilla, 1997
). The negative region of protein near the extracellular S4 segment at
90 mV is consistent with the pH data, and the positive region of protein at
90 mV is consistent with the iodide quenching data. The relative proximity of protein
regions to the residues reflects the anisotropy data.
This vestibule may also be involved in inhibiting the
movement of the voltage sensor after the attachment of
charged methanethiosulfonate reagents to sites M356C,
A359C, and V363C near the S4 segment (Cheney et al.,
1998
). Although this gives no more than a rough picture of the environment surrounding the extracellular
portion of the S4 segment, it gives a reasonable physical
basis to explain the quenching of the fluorophore by the lining of the hydrophilic vestibule.
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FOOTNOTES |
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Address correspondence to Dr. F. Bezanilla, Dept. of Physiology, UCLA School of Medicine, 10833 Le Conte Avenue, Los Angeles, CA 90095. Fax: 310-794-9612; E-mail: fbezanil{at}ucla.edu
Original version received 19 May 1998 and accepted version received 28 July 1998.
We thank Dr. Adrian Gross for his help in editing the manuscript. Special thanks go to Dr. Paul Selvin for his contributions and insight into the polarization and anisotropy studies.
This work was supported by National Institutes of Health grant GM-30376 and the Hagiwara Chair funds to F. Bezanilla. A. Cha is also supported by the UCLA Medical Scientist Training Program (GM-08042) and a National Research Service Award from the National Institute of Mental Health (MH-12087).
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Abbreviations used in this paper |
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dF, fluorescence difference; R, intensity ratio; TEA, tetraethylammonium; TMRM, tetramethylrhodamine maleimide.
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