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* Department of Pharmacology and the Biomedical Engineering Program, University of Nevada, Reno, Nevada; and Department of Physiology and Biophysics, University of Calgary, Calgary, Alberta T2N 4N1, Canada
Correspondence: Address reprint requests to John Sutko, Dept. of Pharmacology, 318 Howard Bldg., Room 214, University of Nevada, Reno, NV 89557. Tel.: 775-784-4121; Fax: 775-784-1620; E-mail: sutko{at}unr.edu.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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), has provided a noninvasive experimental approach for assessing RyR channel function in intact cells. At the protein level, site-directed incorporation of fluorescent reporter molecules has been used to probe changes in the structure of transporters, such as the lactose permease in Escherichia coli (Vazquez-Ibar et al., 2002), and ion channels, such as the Shaker K+ channel (Cha et al., 1999; Sonnleitner et al., 2002), that underlie the function of these proteins.
Our goal was to develop an experimental system for applying optical imaging techniques to investigations of RyR channel properties in vitro. As a consequence of their localization to intracellular membranes, RyR channels have been studied after reconstitution of membrane vesicles or purified protein in planar lipid bilayers (Fill and Copello, 2002; Sitsapesan and Williams, 1990; Smith et al., 1986; Williams et al., 2001). To take advantage of the ability of the bilayer system to measure Ca2+ flux through single RyR channels in a very accurate manner, we sought to develop an optical bilayer system permitting simultaneous imaging of RyR channel-related functions and recording of single channel currents.
Our work was motivated by several experimental objectives. First, analyses of events involved in activation of muscle cells and, in particular, of RyR channel function in intact cells require quantitative information about the Ca2+ flux underlying the imaged Ca2+ spark. However, extraction of this information is complicated by factors and conditions extrinsic to the RyR channel that prevent the imaged spark waveform from faithfully tracking the waveform of the underlying Ca2+ release flux. These factors include binding of Ca2+ to fixed and mobile buffers, diffusion of free Ca2+ and of Ca2+ bound to buffers and to the Ca2+ indicator, and binding of the Ca2+ indicator to cellular proteins. There is also uncertainty about the distance of the plane of focus from sites of Ca2+ release during imaging, and the affinity of the indicator for Ca2+. The kinetics of the indicator-Ca2+ binding reaction may also distort the spark signal. Influences by these factors alter the response by the Ca2+ indicator to the Ca2+ flux waveform in both time and space. Defining the influences of many of these extrinsic factors in cells is problematic, because their effects can be interactive and it is difficult to perturb only a single factor in an intact cell in a controlled and systematic manner. To approach this problem, models and simulations have been used to estimate influences exerted by extrinsic factors (Baylor et al., 2002; Cannell and Soeller, 2002; Izu et al., 2001; Jiang et al., 1999; Pratusevich and Balke, 1996; Rios and Brum, 2002; Smith et al., 1998; Sobie et al., 2002). As a complementary approach to this problem, we sought to develop an experimental system in which Ca2+ flux through single RyR channels could be both imaged with fluorescent Ca2+ indicators and recorded electronically, and where the environment of the Ca2+ indicator could be controlled and manipulated. Such an experimental system would permit influences exerted on Ca2+ sparks by extrinsic factors, such as those noted above, to be investigated in a direct and systematic manner.
An optical bilayer system will also permit other aspects of RyR channel function to be studied in novel ways. For example, multiple RyR channels are thought to cluster in functional Ca2+ release units (CRU) (Franzini-Armstrong et al., 1999) that may activate and inactivate in a coupled manner (Copello et al., 2003; Marx et al., 1998, 2001; Ondrias and Mojzisova, 2002; Wang et al., 2002). Consistent with this notion, multiple RyR channels reconstituted in lipid bilayers from SR membranes exhibit coupled gating and both physical interactions between adjacent RyR channels and sequential activation by Ca2+-induced Ca2+ release have been proposed as mechanisms coordinating this gating (Copello et al., 2003; Marx et al., 1998, 2001). Fluorescence imaging may provide a way to distinguish between these possibilities. There is also increasing information concerning the relationship between the primary sequence and the tertiary structure of RyR channels (Liu et al., 2002; Serysheva et al., 1999; Wagenknecht and Samso, 2002). An optical bilayer system capable of analyzing signals from fluorescent reporter molecules incorporated into RyR channels in a site-directed manner will facilitate investigations of the structure and function of this protein similar to those noted above for the lactose permease and K+ channels. The feasibility of this approach is supported by observation of function-dependent fluorescent signals from populations of RyR channels (Ikemoto and Yamamoto, 2002; Ohkusa et al., 1991). As described in this report, as well as in an abstract (Peng et al., 2003), we have developed an optical bilayer system with the sought-after capabilities. As described in the accompanying report (Peng et al., 2004), we have utilized this system to characterize lateral diffusion by single RyR2 channels in bilayers and to identify a mechanism for preventing this movement to enhance the spatial and temporal resolutions of optical imaging related to RyR channel function.
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MATERIALS AND METHODS |
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Preparation of RyR2
Hearts were obtained from adult sheep sacrificed for other purposes at the University of Nevada farm. Sarcoplasmic reticulum membranes were isolated using differential centrifugation and sedimentation through discontinuous sucrose gradients as described previously (Airey et al., 1990; Sitsapesan and Williams, 1994). RyR2 channels were solubilized by incubating SR membranes (3 mg protein/ml) with 2% CHAPS and 1% phosphatidylcholine (PC) in a solution containing (in mM): 500 KCl, 5 DTT 20 Tris (pH 7.4) for 30 min on ice. Nonsolubilized material was removed by centrifugation at 100,000 x g for 30 min. Solubilized RyR channels were purified further by sedimentation through a continuous 10–30% sucrose gradient in a Beckman SW28 rotor at 24,000 x g for 16 h (Airey et al., 1990). Partially purified RyR channels were reconstituted into liposomes by dialysis (8 h x 4 solution changes) against 2 liters of a solution containing, in mM: 250 NaCl, 0.2 CaCl2, 0.1 EGTA, 10 Mops (pH 7.4), 1 DTT. After dialysis, liposomes were pelleted by centrifugation at 266,000 x g for 50 min in a Beckman TL-100 rotor, resuspended in a solution containing, in mM: 500 KCl, 20 Tris/Hepes (pH 7.4), 400 sucrose, 5 DTT, rapidly frozen in liquid N2 and stored at -80°C. All solutions contained the following protease inhibitors: phenylmethylsulfonyl fluoride (0.23 mM), soybean trypsin inhibitor (10 µg/ml), aprotinin (1.0 µg/ml), pepstatin A (1.0 µM), benzamidine (0.2 mM), and leupeptin (1.0 µM).
The optical bilayer system
The optical bilayer system used in the studies described in this and the accompanying report is schematized in Fig. 1. A small trans chamber was attached via a glass capillary tube to the head stage of an Axopatch 1-B patch clamp amplifier (Axon Instruments, Union City, CA) using a microelectrode holder (E.W. Wright, Guilford, CT). The head stage was, in turn, mounted on a 3-axis hydraulic manipulator (Narashige, East Meadow, NY), which was used to position the trans chamber within the cis chamber. A water immersion microscope objective (70x, 1.2 NA, Lomo Optics, Germantown, MD) was introduced in a horizontal orientation through the front wall of the cis chamber. Focusing was achieved by using a computer-driven stepper motor to adjust the position of the trans chamber relative to the objective. A small magnetic stir motor (not shown) was mounted in the rear wall of the cis chamber, where it held a small stir bar used to mix the contents of the cis chamber.
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160 µm), exhibits a steep profile. As a consequence, the portion of the bilayer that can be scanned without interruption of the beam by the edges of the front wall of the trans chamber decreases rapidly with distance from the front of the chamber. An estimate of the maximal depth compatible with uninterrupted scanning of a bilayer with our optical system is shown in panel B of Fig. 2. A fluorescent solution was placed in both the cis and trans chambers in the absence of a bilayer. As the plane of focus was advanced into the hole that normally contained the bilayer the imaged fluorescence intensity remained constant until a depth of 18 µm was reached. Beyond this point the fluorescence intensity declined rapidly indicating that the volume of fluorophore-containing solution being sampled was diminished. Since bilayers frequently form as far away from the front surface of the trans chamber as possible, it was necessary to use holes <15 µm deep. To meet this requirement, holes 50–100 µm in diameter were formed in small rectangular pieces of 12.5 µm thick Halar film. As shown in Fig. 3, the entire bilayer can be scanned when holes limited to this depth are used. The bilayer was imaged using reflected red (633 nm) light and a fluo-3 excitation/emission filter/dichroic beam splitter set. This arrangement permits imaging of the bilayer surface, as well as the ability to determine the distance between the bilayer and the front surface of the trans chamber. The latter information is necessary for analysis of how variations in the distance between the plane of focus and the plane of the RyR channel influence spark properties.
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1 s to acquire. The x-t line scans shown in Figs. 4 and 8 contain 512 lines and again required 1 s to acquire.
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), suitable for activating RyR2 channels. The trans chamber solution contained 50 mM CaCl2 and 10 mM Tris/Hepes, pH 7.4. In the experiments described in Figs. 8 and 9, 2 µM annexin 12 was added to the trans chamber to restrict lateral diffusion of the RyR2 channel (see Peng et al., 2004, for details). Liposomes containing RyR2 channels were added to the cis chamber and fusion with the bilayer was enhanced by creation of an osmotic gradient by addition of 3M KCl to the cis chamber. After fusion occurred, the cis chamber was perfused while stirring with 5 vol of a solution containing 85 mM Tris/Hepes, pH 7.4. When perfusion was complete a final concentration of 25–75 µM fluo-3 was added to the cis chamber. Trans-bilayer currents were recorded and analyzed using a system comprised of an Axopatch 1B patch clamp amplifier, a Digidata 1200 interface, a PC computer, and pClamp 8 software.
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) >99% of the trans-cis current will be carried by Ca2+, as calculated using the model described by Tinker et al. (1993) (Alan Williams, personal communication).
A voltage command (VC) protocol was developed that permitted Ca2+ flux through RyR2 channels to be determined solely by the voltage imposed across the bilayer. Using this protocol, the amplitude, time course, shape, and consequently the magnitude, of the Ca2+ flux can be controlled with precision. To achieve such control, the open probability (Po) of RyR2 channels must be maintained at, or close to, 1.0 during the voltage step, so Ca2+ fluxes are not affected by channel gating. To keep background fluo-3 fluorescence at acceptably low levels, it was also necessary to achieve this level of activation in the presence of submicromolar concentrations of free Ca2+ in the cis chamber. Based on the findings of Sitsapesan and Williams (1996) and Sitsapesan (1999) these requirements were met by addition of suramin (100–200 µM) to the cis chamber in the presence of 0.1 µM free Ca2+. To prevent leak of Ca2+ through suramin-activated RyR2 channels, the trans-bilayer voltage was maintained at the reversal potential for RyR2 channel-mediated Ca2+ currents, typically -60 mV (trans relative to cis) between measurements. A comparable reversal potential was calculated using the model of Tinker et al. (1993) (Alan Williams, personal communication). The voltage was stepped to more positive trans voltages to elicit a net trans-to-cis Ca2+ flux. Using the VC protocol, reproducible Ca2+ fluxes and associated fluo-3 fluorescence responses could be achieved in a repetitive manner. Negative controls demonstrating the absence of fluo-3 fluorescence when there was no trans-to-cis Ca2+ current were obtained 1), from records where channels either failed to open at all or closed for a significant period during a step to positive voltages; 2), when the applied voltage was maintained at the reversal potential; and 3), when steps to voltages more negative than the holding potential were applied.
Isolation of ventricular cardiomyocytes from adult mice
Myocytes were isolated using standard techniques (Duan et al., 1999), permitted to recover to a Ca2+-tolerant state, and loaded with the Ca2+ indicator, fluo-3 AM, using methods described by Ritter and co-workers (2000). Myocytes were permitted to settle onto cover slips coated with either polylysine or laminin, and the coverslips were fixed to the front wall of the trans chamber in an orientation identical to that of the lipid bilayer. Fluorescence imaging was then performed utilizing the same illumination, optics, and detection system used when a bilayer was present. In some experiments extracellular K+ was elevated to 10-20 mM to increase the frequency of spontaneously occurring sparks.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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) and comparable to those observed for sparks imaged in cardiomyocytes from other species (see compilation in Table 1 of Rios and Brum, 2002).
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30 ms and then reopened while the position of the channel was being scanned. Because the x,y scan was acquired left to right from top to bottom, this resulted in bifurcation of the fluorescence signal. An intensity-time trace obtained from top to bottom through the center of the spark (white vertical line in panel C) is aligned with the current trace in panel D. The sharp demarcation of the period of channel closure, as well as the ramp responses shown in Fig. 8, and the x-t images shown in Fig. 5 in Peng et al. (2004) illustrate the kinetic properties of the optical system to changes in Ca2+ in the vicinity of the bilayer.
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0.5–1.4 pA (Mejia-Alvarez et al., 1999; Tinker et al., 1993). These studies used luminal Ca2+ concentrations of 1–10 mM. With these estimates, the range of RyR2 channel current amplitude in Fig. 7 is equivalent to the flux of Ca2+ through 1 to >20 RyR2 channels. Ca2+ flux is expressed as peak amplitude rather than as the total or cumulative net flux of Ca2+ obtained by integrating the current amplitude over the duration of the voltage step in this figure. Since the same duration was used for all voltage steps and Po 1.0, integration resulted in only a proportional shift of the values and did not alter qualitative aspects of the relationship between fluorescence and Ca2+ flux. As noted in the Introduction, RyR2 channels in a CRU may activate in a coordinated manner. In this case, RyR2 channels mediating the Ca2+ flux responsible for a Ca2+ spark could activate and inactivate at the same time and this type of activity could be mimicked as a square step in RyR2 channel Ca2+ current, similar to that shown in Fig. 5 of the accompanying report. Alternatively, individual RyR2 channels could activate and inactivate at different rates and a more complex current waveform would be required to simulate this type of activity. For example, fluo-3 fluorescence elicited by ramping rather than stepping the trans-bilayer voltage is shown in Fig. 8. The ramp protocol produces a rise and fall in fluo-3 fluorescence, which in some cases may be more similar to the rising phase of sparks imaged in cardiomyocytes. The ability to elicit arbitrarily complex Ca2+ flux waveforms demonstrates further the ability of the VC protocol to mimic aspects of the relationships between optical recordings and the flux of Ca2+ through RyR2 channels in cells. The change in fluorescence intensity shown in this figure is in good agreement temporally with the imposed change in voltage, again illustrating the temporal properties of the optical bilayer system.
To simplify initial studies, purified RyR2 channels were reconstituted in liposomes at a low protein/lipid ratio to enhance incorporation of single channels. However, on occasion RyR2 channel Ca2+ currents that were multiples of the unitary current were recorded indicating the presence of more than one active RyR2 channel in the bilayer. As shown in the x,y images presented in Fig. 9, when this occurred multiple foci of fluo-3 fluorescence were imaged. In the experiment shown in the upper left panel of the figure, one of two RyR2 channels did not open until its position in the bilayer had already been scanned. Consequently, only a single fluorescence signal was observed. During the next scan shown in the lower left panel, both RyR2 channels activated and two fluorescence signals were imaged. Three-dimensional representations of the x,y images are presented in the panels on the right side of the figure. Inspection of the intensity values indicated that the flattened tops of the signals are due to the square shape of the fluorescence signal and not to signal saturation. Another example of this shape can be seen in the intensity-time traces shown in Fig. 5 in Peng et al. (2004). These data demonstrate further the ability to establish the location of active RyR channels using the optical bilayer system.
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DISCUSSION |
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, we describe use of this system to characterize lateral diffusion of RyR2 channels in bilayers and identification of the ability of annexin 12 to effectively immobilize RyR2 channels within the bilayer. Below, we discuss additional technical aspects of the bilayer system and application of this system to three areas of current research.
Additional considerations concerning the optical bilayer system
The optical bilayer system can be used with both widefield (nonscanned) and laser scanning confocal fluorescence imaging systems (LSCS). We implemented a LSCS for three reasons. First, as discussed below, a motivation for developing an optical bilayer system was to provide a complementary approach for characterizing the relationships between Ca2+ indicator fluorescence and the flux of Ca2+ through RyR2 channels. LSCS have been used in the majority of studies in the literature that describe properties of Ca2+ sparks imaged in cells. Therefore, we felt it was necessary to use a similar imaging system to make meaningful comparisons between reported data and results obtained with the bilayer system.
Second, LSCS image fluorescence in a restricted volume. This ability is important for the temporal resolution observed for fluorescence imaged in the optical bilayer system. As shown in Figs. 4 and 9 of this report and Fig. 5 of Peng et al. (2004), the temporal and dynamic resolution of fluo-3 fluorescence tracks elicited Ca2+ fluxes in a reasonable manner. The rapidity of the diffusion of Ca2+ from the restricted volume being imaged is one reason for the temporal resolution achievable with the optical bilayer system. Future studies will address the extent to which the on- and off-rates of the indicator-Ca2+ binding reaction and rebinding of Ca2+ during a single Ca2+ flux event contribute to the time course observed for the fluorescent responses.
A third reason for use of a LSCS was to utilize detectors with the greatest data acquisition rates. Currently, single point detectors, such as photodiodes and photomultiplier tubes, have greater sampling rates than detector arrays, such as CCD cameras, typically used with nonscanned widefield systems. The ability to immobilize RyR2 channels in the bilayer described in the accompanying report will permit parking the laser beam on the position of the channel and recording fluorescence with photon-limited temporal resolution.
A potential complicating factor associated with the optical bilayer system is that under certain conditions the structure and properties of the bilayer may influence fluorescence responses to Ca2+ fluxes. Several factors may create asymmetry in the fluorescent signal. These include an illusion of asymmetry when a channel is located at the edge of the bilayer, resulting in greater intensity values on one side of the channel, particularly as background Ca2+ levels are increased by RyR2 channel activity. For example, this can be appreciated by comparing the bilayer fluorescence signals shown in panels A and B in Fig. 5. Second, curvature of the bilayer as it approaches the torus can produce spatial aberrations. Third, care must be taken to ensure that the face of the objective is parallel with the plane of the bilayer. Properties of the optical system may also contribute to signal asymmetry. As bearing surfaces in scan mirror galvanometers age, they can become nonlinear and result in distorted spatial representations. Also, depending on their design, microscope objectives have varying degrees of spherical aberration.
Application of the optical bilayer system to areas of current research
Analysis of Ca2+ sparks
Ca2+ sparks offer unique insights into cellular activation and the function of RyR2 Ca2+ release channels in intact cells. A rigorous analysis of RyR2 channel activity in situ requires accurate estimates of the magnitude of the Ca2+ flux through single RyR2 channels, the total quantity or mass of Ca2+ underlying a spark, and the temporal properties of the Ca2+ flux. However these estimates are difficult to make, since the waveform of the spark is distorted from that of the underlying Ca2+ flux by cellular factors extrinsic to RyR2 channels. These factors include concerted gating of multiple RyR2 channels, diffusion of free Ca2+ and Ca2+ bound to mobile buffers, the kinetics of Ca2+ binding to the Ca2+ indicator, the extent of binding of the Ca2+ indicator to cellular proteins, release of Ca2+ into a restricted space, competition for Ca2+ with removal processes, and having the plane of focus distant from the site of Ca2+ release. The optical bilayer system can be used to compare cellular sparks with bilayer sparks obtained in response to a known range of Ca2+ fluxes elicited with the VC protocol that encompass those thought to occur in intact cardiomyocytes. As shown in Fig. 4, it is possible to compare bilayer sparks with cellular Ca2+ sparks imaged with the same optical system.
Two approaches have been used to measure the Ca2+ flux underlying sparks imaged in cardiac myocytes. In technically elegant experiments, Wang et al. (2001) used fluo-4 fluorescence responses to Ca2+ fluxes through voltage-gated L-type Ca2+ channels to calibrate the Ca2+ flux underlying Ca2+ sparks measured in the same rat ventricular myocyte and estimated that a spark involves 2.1 pA of Ca2+ flux (also see Cheng and Wang, 2002). As noted in the Introduction, others have used simulation and modeling to account for influences of factors, such as those discussed in the preceding paragraph, and estimate the Ca2+ flux underlying a spark. In general, estimates of Ca2+ fluxes associated with sparks obtained by modeling exceed that of Wang et al. (2001). For example, Soeller and Cannell (2002) concluded that sparks imaged in rat ventricular cardiomyocytes involve a Ca flux >7.5 pA. Both imaging and modeling approaches should yield similar estimates of the Ca2+ flux underlying a spark, and as noted by Rios and Brum (2002), discrepancies between these estimates may in part be due to systematic selection of different size sparks by the different laboratories. However, Baylor et al. (2002) demonstrated recently that the accuracy of simulations of Ca2+ sparks imaged in frog skeletal muscle fibers is greatly enhanced by improved estimates of Ca2+ binding to cellular proteins and the properties of Ca2+ binding to the Ca2+ indicator, fluo-3, and by maximizing the spatial resolution of the optical system. This work indicates the importance of a more complete understanding of the effects of cellular factors on the spatial and temporal properties of Ca2+ sparks. Consequently, a rationale for development of the optical bilayer was to provide a system suitable for a bottom-up assessment of their impact on the properties of a "spark" waveform in a direct and systematic manner.
The VC protocol is an important component in our approach. Setting the Po of a RyR2 channel to 1.0 permits precise control over the amplitude, time course, and shape of Ca2+ fluxes through single RyR2 channels. Since RyR2 channel currents are measured, any occasional deviations from a unitary Po are taken into account. The ability to control Ca2+ flux allows systematic variation of its magnitude and time course and ranges of values for parameters of RyR channel function to be investigated. This avoids bias and assumptions about the values of RyR2 channel parameters in intact cells, where little is known with certainty. Ca2+ sparks are generally accepted to be the product of Ca2+ flux through multiple RyR2 channels localized in a CRU. Using the estimates of unitary physiological RyR2 channel Ca2+ conductance by Tinker et al. (1993) and Mejia-Alvarez et al. (1999), the range of RyR2 channel Ca2+ current amplitudes achievable with saturating levels of trans Ca2+ in the optical bilayer system is sufficient to mimic unitary Ca2+ currents through 1 to >20 RyR2 channels. Moreover, as shown in Fig. 9, voltage ramps can be used to vary the steepness and shape of the rising and falling phases of the command signal to mimic nonhomogeneities in the rates of activation and/or inactivation of individual RyR2 channels within a CRU. The ability to immobilize RyR2 channels in bilayers described in Peng et al. (2004) makes it possible to use x-t or line scans through the position of the functional RyR2 channel for repetitive analysis of Ca2+ indicator responses to measured Ca2+ fluxes. The reproducible nature of the fluorescence signals to consecutive voltage steps (see Fig. 8 in this report, and Fig. 5 of Peng et al., 2004) make it possible to use signal averaging to improve signal-to-noise.
Coupled gating of RyR channels
Analysis of the mechanism(s) underlying coupled gating of RyR channels may also benefit from application of the optical bilayer system. Studies by Marx et al. (1998, 2001) indicate that multiple RyR channels in bilayers can exhibit coupled gating. Exposure to rapamycin, which causes FK506 binding proteins (FKBP) to dissociate from RyR channels, resulted in non-coordinated gating and it was suggested that coordination of gating may involve interactions between adjacent RyR channels stabilized by binding of FKBP to RyR. Activation of RyR channels by the Ca2+-induced Ca2+ release (CICR) mechanism (Fabiato and Fabiato, 1975) could provide an alternative or additional mechanism for coordinating activation of closely grouped RyR channels (Copello et al., 2003). In this case, Ca2+ released from one RyR channel would activate one or more nearby RyR channels in a sequential or parallel manner. The CICR mechanism would not require physical contact between RyR and may permit coordination of RyR channels separated by greater distances. The optical bilayer system may help to distinguish between these mechanisms. If distances between RyR channels coordinated by CICR are sufficiently great, it will be possible to resolve Ca2+ indicator signals having multiple peak amplitudes. The centroid of each peak can be determined to yield estimates of inter-RyR channel distances. Since the molecular dimensions of RyR channels are known (Wagenknecht and Samso, 2002), it should be possible to decide whether these distances are compatible with physical contact between RyR channels.
Structure-function relationships of RyR channels
A long-term goal for development of an optical bilayer system is to provide one of the tools required for analyses of the changes in RyR channel structure responsible for channel function. Significant steps have been made toward providing the other tools required for such experiments. As discussed in the Introduction, the relationship between the primary sequence and the tertiary structure of RyR channels is being elucidated using cryoelectron microscopy. Although not capable of atomic resolution, this approach does provide an initial basis set of information. In addition, significant advances have been made in producing diffracting crystals of membrane proteins (Jiang et al., 2002, 2003; Nishida and MacKinnon, 2002). Studies of RyR channel structure-function relationships will be facilitated by the use of molecular modeling based on structural homology with other ion channels (Williams et al., 2001). New strategies for introducing reporter groups with conformation-dependent fluorescence signals into RyR channels in a site-directed manner will have to be developed; however, work by Ikemoto and coworkers who have recorded variations in fluorescence intensity from populations of labeled RyR channels during release of Ca2+ from SR vesicles (Ohkusa et al., 1991; Yamamoto and Ikemoto, 2002) supports the feasibility of this step. Finally, recent studies involving K+ and gramicidin channels demonstrate the feasibility of recording channel conformation-dependent fluorescence signals from single ion channels (Borisenko et al., 2003; Sonnleitner et al., 2002).
The abilities to image fluorescence responses to single RyR2 channel Ca2+ currents and to immobilize RyR channels in bilayers described for the first time in this and the accompanying report will make important contributions to studies of single RyR channels. As noted in a preliminary report, not all fluorescently labeled channel proteins that associate with a bilayer reconstitute as functional ion channels (Blunck et al., 2003). This makes it necessary to distinguish fluorescence signals associated with active channels. This situation is confounded by the fact that many fluorophores emit light in an intermittent manner, which could be interpreted as a functionally related signal, but be derived from an inactive channel. To avoid this problem, it is essential to identify the ion channel(s) conducting the currents being recorded. The ability to image Ca2+ flux through single (or multiple) RyR channels demonstrated in this report provides a way to precisely locate the position of active RyR channel(s) in the bilayer.
In conclusion, we have developed an optical bilayer system capable of simultaneous imaging of Ca2+ flux through single RyR2 channels reconstituted in planar lipid bilayers and electrical recording of single channel currents. Capabilities of this system are demonstrated and its utility for addressing several current research problems is discussed.
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ACKNOWLEDGEMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGEMENTSREFERENCES |
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The authors of this report received support from the following sources: J.A.A., S.P., and J.L.S. from National Institutes of Health grants HL53677 and AR45112; N.G.P. and J.L.S. from a National Science Foundation EPSCoR grant EPS0132556: G.J.K. from the Canadian Institutes of Health Research and the Heart and Stroke Foundation of Alberta; and D.D. from National Institutes of Health grants HL63914 and NCRR P20 RR15581.
Submitted on July 18, 2003; accepted for publication September 4, 2003.
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