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Am J Physiol Cell Physiol 284: C1654-C1668, 2003. First published February 12, 2003; doi:10.1152/ajpcell.00469.2002
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Vol. 284, Issue 6, C1654-C1668, June 2003

Two-photon molecular excitation imaging of Ca2+ transients in Langendorff-perfused mouse hearts

Michael Rubart1, Exing Wang2, Kenneth W. Dunn2, and Loren J. Field1

1 Wells Center for Pediatric Research and Krannert Institute of Cardiology, and 2 Department of Medicine, Division of Nephrology, Indiana University School of Medicine, Indianapolis, Indiana 46202


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The ability to image calcium signals at subcellular levels within the intact depolarizing heart could provide valuable information toward a more integrated understanding of cardiac function. Accordingly, a system combining two-photon excitation with laser-scanning microscopy was developed to monitor electrically evoked [Ca2+]i transients in individual cardiomyocytes within noncontracting Langendorff-perfused mouse hearts. [Ca2+]i transients were recorded at depths <= 100 µm from the epicardial surface with the fluorescent indicators rhod-2 or fura-2 in the presence of the excitation-contraction uncoupler cytochalasin D. Evoked [Ca2+]i transients were highly synchronized among neighboring cardiomyocytes. At 1 Hz, the times from 90 to 50% (t90-50%) and from 50 to 10% (t50-10%) of the peak [Ca2+]i were (means ± SE) 73 ± 4 and 126 ± 10 ms, respectively, and at 2 Hz, 62 ± 3 and 94 ± 6 ms (n = 19, P < 0.05 vs. 1 Hz) in rhod-2-loaded cardiomyocytes. [Ca2+]i decay was markedly slower in fura-2-loaded hearts (t90-50% at 1 Hz, 128 ± 9 ms and at 2 Hz, 88 ± 5 ms; t50-10% at 1 Hz, 214 ± 18 ms and at 2 Hz, 163 ± 7 ms; n = 19, P < 0.05 vs. rhod-2). Fura-2-induced deceleration of [Ca2+]i decline resulted from increased cytosolic Ca2+ buffering, because the kinetics of rhod-2 decay resembled those obtained with fura-2 after incorporation of the Ca2+ chelator BAPTA. Propagating calcium waves and [Ca2+]i amplitude alternans were readily detected in paced hearts. This approach should be of general utility to monitor the consequences of genetic and/or functional heterogeneity in cellular calcium signaling within whole mouse hearts at tissue depths that have been inaccessible to single-photon imaging.

rhod-2; fura-2; BAPTA; 2,3-butanedione monoxime; cytochalasin D


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

TWO-PHOTON MOLECULAR EXCITATION (TPME) fluorescence microscopy offers advantages over traditional confocal approaches in that it permits the acquisition of fluorescent signals originating deep within living, strongly light-scattering tissues (11, 13, 15, 29, 39, 49, 57). This is accomplished by exciting the sample with high-intensity laser light at a wavelength approximately double what is normally required to excite the fluorophore. By rapidly pulsing the laser light, an extremely high photon density localized to the diffraction-limited volume of the objective lens focal point is achieved. Because lower energy photons are used, fluorophore emission occurs only following excitation by two or more photons. Because the probability of achieving two-photon excitation declines rapidly with the fourth power of distance from the focal point, excitation (and thus emission) is confined to an extremely thin optical section. Consequently, all of the fluorescent signal can be collected to generate the image. Because TPME imaging does not suffer from degradation of signal-to-background ratio to nearly the same extent as confocal imaging, it provides a high-contrast image even at significant depths in strongly scattering tissue (13). This property was illustrated by Yuste and Denk (57), who were able to resolve calcium transients in vivo at the level of single dendritic spines projecting to a depth of up to 150 µm within the cortical surface of the brain. On the basis of these collective properties, TPME might provide a powerful methodology with which to image calcium transients in individual cardiomyocytes within intact hearts.

Although single-photon laser scanning microscopy has been used extensively to study subcellular Ca2+ signals in isolated cardiomyocytes (12, 30), there is only a limited number of studies wherein individual cells within intact cardiac tissue were imaged. For example, Wier and coworkers (54) developed a system for imaging subcellular calcium levels in isolated, superfused rat papillary muscles. The muscle preparations were loaded with the calcium fluorophore fluo-3 using iontophoresis, and both calcium sparks and calcium waves were imaged at depths up to 40 µm from the endocardial surface. More recently, Minamikawa and colleagues (32) recorded calcium transients and calcium waves in single cardiomyocytes within the superficial epicardial layer of Langendorff-perfused rat hearts using fluo-3. Finally, Kaneko and coworkers (27) were able to assess the detailed quantitative properties of sporadic calcium waves in intact rat hearts using real-time confocal microscopy and fluo-3. However, no analyses of the quantitative properties of electrically evoked calcium transients were performed.

The ability to image calcium signals at subcellular levels within the intact heart is an important goal, because it could provide valuable information toward a more integrated and comprehensive understanding of calcium regulation in cardiac muscle (7). To accomplish this, TPME in combination with scanning microscopy was employed to measure [Ca2+]i-dependent changes in fluorescence intensity of the calcium indicators rhod-2 and fura-2 at the single cardiomyocyte level in a buffer-perfused mouse heart preparation in the presence of the excitation-contraction uncoupler cytochalasin D (8). This system should exploit the anticipated benefits of TPME (namely improved signal-to-background ratio even at significant tissue depths without loss of spatial discrimination). With the use of this approach, it is possible to examine electrically evoked calcium transients at depths <= 100 µm below the epicardial surface and also to derive physiologically meaningful information about amplitude and kinetics of [Ca2+]i transients. Comparative analyses indicated that, under the conditions employed, rhod-2 and cytochalasin D are superior reagents for monitoring [Ca2+]i transients and effecting excitation-contraction uncoupling. The potential utility of the system is further demonstrated by studies in normal and transgenic tissues to identify physiological (i.e., calcium waves) and pathophysiological (i.e., calcium transient amplitude alternans) abnormalities in calcium handling in the paced heart. This is, to the best of our knowledge, the first methodology that permits visualization of electrical stimulus-induced [Ca2+]i transients at the single-cell level within the whole heart. The potential usefulness of this system for functional assessment of the heart is discussed.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Heart preparations. Adult [C3Heb/FeJ × DBA/2J] F1 mice were used for all studies (inbred progenitors were obtained from the Jackson Laboratory, Bar Harbor, ME). Fifteen minutes after intraperitoneal injection of heparin (125 IU/kg body wt), the heart was excised, the ascending aorta was cannulated with a customized no. 18 hypodermic needle (length: 1 in.), and hearts were perfused in the Langendorff mode. Perfusion was carried out at a constant mean perfusion pressure of 60 cmH2O and at 21°C with oxygenated (100% O2) Tyrode's solution containing (in mmol/l) 134 NaCl, 4 KCl, 1.2 MgSO4, 1.2 NaH2PO4, 10 HEPES, 11 D-glucose, and 2 CaCl2 (pH = 7.35 adjusted with 1 mol/l NaOH). During dye loading and washing out, the solution also contained 50 mmol/l butane dione monoxime (BDM). During [Ca2+]i imaging, cytochalasin D (50 µmol/l; stock solution: 3.9 mmol/l in DMSO) was added to the Tyrode's solution to eliminate contraction-induced movement of the heart (8). Preliminary experiments showed that spontaneous heart rates could exceed 400 beats/min under the experimental conditions employed. Because the limited acquisition time of our scanning microscope in line scan mode (maximal scan speed: 32 ms/line) precludes undistorted examination of [Ca2+]i transients at such high rates, the heart rate was lowered to <60 beats/min by adding acetylcholine to the perfusate (10 µmol/l; stock solution: 10 mmol/l in deionized water).

After an initial perfusion period of ~30 min, the buffer was switched to Tyrode's solution containing the acetoxymethylesters (AM) of the calcium fluorophores rhod-2 or fura-2 (10 µmol/l; Molecular Probes, Eugene, OR). The rhod-2/AM solution was prepared by dissolving 168 µg of the indicator in 150 µl of DMSO, mixing it with 17 µl of Pluronic F-127 (25% wt/vol) for 2 min, and finally diluting the mixture in 15 ml of Tyrode's solution. Aliquots of a 1-mmol/l fura-2/AM stock solution (solvent: dry DMSO) were added to 15 ml of Tyrode's solution to obtain a final concentration of 10 µmol/l. After a 15-min loading period, the heart was perfused with dye-free Tyrode's solution for 20 min to allow for deesterification of the dyes by endogenous esterases. Deesterified rhod-2 and fura-2 were retained within the cells, permitting imaging of [Ca2+]i. In contrast, rhod-2/AM and fura-2/AM were washed out. The plasma membrane-selective fluorescent indicator di-4-ANEPPS (Molecular Probes) was administered in a bolus (3 µg) through a site port in the perfusion system.

For two-photon imaging, the perfused heart was placed in a circular dish (23-mm diameter; Bioptechs, Butler, PA) with the anterior left ventricular epicardial surface down. To optimize the focal plane, the heart was gently pressed against the coverslip (170-µm thickness) at the bottom of the chamber by means of a bipolar platinum stimulation electrode (interelectrode distance: 0.5 mm) placed on the right epicardial surface. Hearts were stimulated at 1-2 Hz via 2-ms square wave pulses with ×1.5 threshold current amplitude. The stimuli were delivered by a constant-current isolator (Krannert Engineering, Indianapolis, IN) driven by a programmable stimulator (SD9, Grass Instruments). With the use of these stimulation parameters, it was shown in preliminary experiments that the average width of stimulated QRS complexes in volume-conducted electrocardiograms (recorded from a Langendorff-perfused mouse heart) was greater than that of QRS complexes during spontaneous sinus rhythm, indicating that the hearts were effectively paced rather than field stimulated.

TPME imaging. Images were recorded with a Bio-Rad MRC 1024 laser scanning microscope, which was modified for TPME. Illumination for TPME was provided by a mode-locked Ti:Sapphire laser (tuning range: 740-890 nm; Spectraphysics, Mountain View, CA), which generated a train of 100-fs pulses at a repetition rate of 82 MHz, which is significantly longer than the mean fluorophore excited-state lifetime of typically 1 to 2 × 10-9 s (17, 38). The laser was tuned to a center wavelength of 810 nm for excitation of rhod-2, calcein, and di-4-ANEPPS and 800 nm for fura-2. We selected the wavelength for fura-2 excitation on the basis of measurements of two-photon action cross sections previously published by Xu et al. (56). Preliminary measurements of rhod-2 emission in a rhod-2-loaded heart at wavelengths ranging from 750 to 860 nm showed a modest peak at a wavelength of 810 nm. We subsequently used this wavelength for all rhod-2 experiments. The heart was imaged through a Plan Apo Nikon ×60 1.2 numerical aperture water-immersion lens. Energy of the laser light was adjusted such that no saturation occurred. The emitted light was split by two dichroic mirrors (550-nm long pass and 500-nm short pass) in series, passed through narrow band-pass filters (560-650 and 500-550 nm), and collected with external photomultiplier tubes (PMT). Thus emission from the fluorophore was not descanned using a pinhole as in confocal microscopy. Images at each focal plane were collected at a resolution of 0.43 × 0.43 µm/pixel along the x/y-axis. The intensity of each pixel was digitized at 8-bit depth and stored on the computer's hard disk for off-line analysis.

Intracellular calcium transients were recorded in the full-frame and line-scan mode of the microscope. To generate full-frame images (512 × 512 pixels), fluorescence signals scanned on horizontal (x, y) planes were digitized and stored directly on the hard disk. Pixel size, as set by the objective and hardware zoom factor of the laser scanning unit, ranged from 0.108 to 0.43 µm. Scan speed in full-frame mode was 1.33 ms/line. In the line-scan mode, tissue within the focal plane of the objective was scanned at a rate of 31.25 Hz along a line spanning at least two juxtaposed cardiomyocytes (see Fig. 1D). This rate corresponds to a scan speed of 32 ms/line, which is the maximal speed that can be achieved in the line-scan mode of our imaging system. The temporally sequential line scans were then stacked vertically to generate the composite line-scan images. Thus the vertical dimension is time, increasing from top to bottom, and the horizontal dimension is distance along the scan line. The fluorescent profile of [Ca2+]i transients in adjacent myocytes was obtained by averaging the line-plot data of line-scan images from adjoining scan lines around the middle of each cell. Postacquisitional analysis was performed using MetaMorph software version 4.6r5 (Universal Imaging, Downingtown, PA). For determination of the amplitude and time course of stimulation-evoked rhod-2 and fura-2 fluorescence transients, F/Fo ratios and their t90-50% and t50-10% values were calculated, where F is fluorescence intensity, Fo is prestimulus fluorescence intensity, and t90-50% and t50-10% represent the time intervals from 90 to 50% and 50 to 10% relaxation, respectively, measured from the peak of the F/Fo curve. Fo corresponds to the minimal and maximal fluorescence intensity of rhod-2 and fura-2, respectively.


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  		Fig. 1.   Stimulation-evoked single myocyte [Ca2+]i transients in the intact left ventricle of a mouse heart loaded with rhod-2. A: full-frame false color image of a rhod-2-loaded heart obtained during 2-photon excitation at 810 nm. Emission was measured in the 560-650 nm range. White arrows point to endothelial cell nuclei. Scale bar = 20 µm. Scaling was the same in x and y direction. B: rhod-2 intensity profiles from the boxed regions in A. X-axis, distance perpendicular to the scan direction; y-axis, distance along the scan line; z-axis, fluorescence intensity (F) in arbitrary units (a.u.). Numbers 1, 2, and 3 refer to the boxed regions in A. C: full-frame false color image during electrical stimulation at 1 Hz. Image was acquired at a scan rate of 1.33 ms/line during 2-photon excitation at 810 nm, and emission was measured in the 560-650 nm range. A total of 327 lines was imaged. Intensity of Ca2+-bound rhod-2 fluorescence was encoded off-line in shades of red. Simultaneous increases in rhod-2 fluorescence along the length of the horizontal scan line are apparent in the middle of the image, corresponding to a depolarization that occurred when the scan was approximately halfway across the cells in the field of vision. The increase in fluorescence of the myocytes is due to stimulus-evoked increases in [Ca2+]i. White arrowheads point to endothelial cell nuclei. White scale bar = 20 µm. Scaling was the same in x and y direction. D: line-scan image along the red line in C. This single line was scanned 200 times at a rate of 31.25 Hz. The intensity of each line scan was plotted underneath each other to produce the line-scan image. Thus the horizontal dimension is distance along the scan line, whereas the vertical dimension is time (increasing from top to bottom). Horizontal black arrow marks increase in stimulation rate from 1 to 2 Hz. Scale bars = 10 µm horizontally and 700 ms vertically. Black downward arrow indicates direction of time axis. E: time courses of spatially averaged, normalized rhod-2 fluorescence from the regions indicated in D. Tracings were obtained by averaging pixel intensities in the horizontal direction and then dividing the F by the prestimulus F, Fo. The regions were 34, 25, and 23 µm wide, each one representing 1 of 3 juxtaposed cells along the scan line. The numbers in E correspond to the numbers of the line plots identified along the red line in C. F: superimposed tracings of changes in [Ca2+]i as a function of time from the 3 regions indicated in D. Changes in F (Delta F) were normalized by first subtracting the averaged prestimulus F from all data points and then dividing these baseline-corrected values by the peak F. Note the similarity in the time course of the fluorescence ratio from the 3 regions.

For epifluorescence imaging of histological sections, intact hearts were fixed in a solution containing 1% paraformaldehyde, 1.08% cacodylic acid, and 0.67% NaCl; cryoprotected in 30% sucrose; mounted in freezing media; and sectioned at 10 µm. Images were then captured on a Leitz (Solms, Germany) fluorescence microscope using a SPOT digital camera (St. Sterling Heights, MI).

Statistical analysis. Data are reported as means ± SE. ANOVA was used for multiple comparisons. The t-test with the Bonferroni correction was used to identify where the differences among the groups occurred after the significant ANOVA. The paired t-test was used to identify changes within each group. Differences were considered significant at P < 0.05.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Imaging of single myocyte [Ca2+]i transients in intact mouse heart with rhod-2. To image electrical stimulation-evoked [Ca2+]i transients, hearts harvested from adult mice were loaded with rhod-2 via retrograde perfusion on a Langendorff apparatus. The apparatus was then transferred to the microscope stage for imaging. All transients were recorded in the presence of cytochalasin D and acetylcholine to effectively uncouple contraction from excitation (8) and to lower the spontaneous heart rate, respectively. A full-frame TPME image of a rhod-2-loaded heart in the absence of electrical stimulation is shown in Fig. 1A. Intensity profiles from the boxed regions demonstrate that there are no areas of preferred rhod-2 accumulation within the cytosol of cardiomyocytes at rest (Fig. 1B). Small regions of intense rhod-2 fluorescence (white arrows in Fig. 1A) correspond to capillary endothelial cell nuclei. Figure 1C illustrates the distribution of rhod-2 fluorescence during remote electrical stimulation. An increase in relative rhod-2 fluorescence is apparent in the middle of the image, corresponding to a depolarization that occurred when the scan was approximately halfway across the cells in the field of vision (the rate of data acquisition for a full-frame image is slower than the stimulation rate). The rhod-2 fluorescence transient rises simultaneously in all cardiomyocytes, indicating that sarcoplasmic reticulum (SR) calcium release activated by depolarization is highly synchronized over the length of the scan line (even though point stimulation was used). Also, the reduction of rhod-2 fluorescence during the subsequent decline in [Ca2+]i appears to occur synchronously in all muscle cells along the scan line. Because the average conduction velocity in the mouse heart is ~0.5 m/s (3), the time to propagate over the 220 µm sampled by the line scan (440 µs) is still smaller than the line-scan speed (1.33 ms/line). Therefore, propagation delays are not detectable at this time scale.

To determine the time course of changes in [Ca2+]i, calcium transients were also recorded in the line-scan mode (12). The scanned region traversed three juxtaposed cardiomyocytes and is indicated by the horizontal red line in Fig. 1C. An image of the stacked line scans is shown in Fig. 1D, and spatially averaged integrated values for the fluorescence transients from each of the three scanned cardiomyocytes are shown in Fig. 1E. With each stimulus, [Ca2+]i rises rapidly (i.e., within the first pixel) in all cardiomyocytes along the scan line (Fig. 1, D and E), indicating that [Ca2+]i transients are linked to electrical excitation in a 1:1 ratio. The time course of decay of spatially averaged [Ca2+]i shortens with increased pacing rates (Table 1) and occurs with nearly identical kinetics in juxtaposed cardiomyocytes (Fig. 1F). To confirm that the [Ca2+]i transients arise from individual cardiomyocytes, hearts were also loaded with the plasma membrane-selective fluorescent dye di-4-ANEPPS to more clearly delineate cardiomyocyte boundaries during rhod-2 fluorescence imaging. Figure 5 demonstrates that fluorescent signals originating within single cardiomyocyte boundaries were readily imaged.

                              
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  		Table 1.   Properties of electrically evoked rhod-2 and fura-2 transients in single cardiomyocytes within Langendorff-perfused mouse hearts

Numerous previous studies employed the excitation-contraction decoupler BDM to immobilize the heart during fluorescence imaging of changes in transmembrane voltage (4) or intracellular calcium (27). We therefore tested whether this agent could be used as a less expensive alternative to eliminate motion artifacts during TPME imaging. Preliminary experiments showed that BDM at concentrations below 50 mmol/l did not completely suppress visible contractions, whereas higher concentrations resulted in electrical inexcitability as documented by a loss of the stimulated QRS complexes in the volume-conducted ECG. The heart remained inexcitable even after increasing the strength of the electrical stimulus to five times the diastolic threshold. As a consequence, cytosolic calcium transients could not be evoked in response to remote electrical stimulation in the presence of 50 mmol/l BDM as shown in Fig. 2, A and C.


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  		Fig. 2.   Differential effects of 2,3-butanedione monoxime (BDM) and cytochalasin D on excitation-contraction uncoupling. A and B: full-frame false color images during electrical stimulation at 2 Hz in a rhod-2-loaded heart in the presence of 50 mmol/l BDM (A) or 50 µmol/l cytochalasin D (B), respectively. Electrical stimuli were continuously delivered to the right ventricular epicardium during image acquisition. White scale bar = 20 µm. Scaling was the same in x and y direction. C and D: line-scan images along the red lines in A and B. Only sporadic calcium waves occurred in the presence of BDM. No [Ca2+]i transients could be elicited in response to electrical stimulation. In the presence of cytochalasin D, electrical stimuli were associated with [Ca2+]i transients in a 1:1 ratio at either 1 or 2 Hz. White arrows, sporadic calcium waves. Scale bar = 20 µm. Black downward arrow, 2 s.

Sporadic calcium waves, however, continued to occur at this concentration of BDM, indicating that the absence of stimulus-induced calcium transients resulted from a loss of electrical excitability rather than from blockade of the calcium release process. This result is consistent with the previous observations that high concentrations of BDM can inhibit gap junction communication between cardiomyocytes (20, 52). This notion was confirmed by substitution of BDM with cytochalasin D (50 µmol/l) in the perfusate; electrical excitability was restored as documented by the 1:1 ratio of calcium transients and depolarizing stimuli (Fig. 2, B and D). To exclude the possibility that stimuli-induced changes in the rhod-2 signal reflect motion artifacts rather than changes in the intracellular concentration of free calcium, fluorescence of the calcium-insensitive indicator calcein and rhod-2 was simultaneously imaged (Fig. 3). This experiment demonstrates that electrical stimulation only causes changes in the intensity of the rhod-2 signal, whereas the calcein signal remains unaffected by depolarization. Collectively, these results confirm that cytochalasin D eliminates contraction-induced movement of the heart at the microscopic level but does not abolish the stimulus-evoked [Ca2+]i transient. Figure 4A illustrates time courses of spatially averaged, normalized rhod-2 transients obtained from line-scan recordings at depths of 30, 50, and 70 µm. In some cases, [Ca2+]i transients could be measured at depths of up to 100 µm (not shown). Although the relative fluorescence of the electrical stimulation-induced changes in rhod-2 fluorescence varies in the examples shown due to variations in the laser input intensity, the kinetics of the normalized fluorescence transients recorded at various tissue depths are essentially superimposable (Fig. 4B).


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  		Fig. 3.   Absence of motion artifacts in cytochalasin D-treated hearts. A: full-frame pseudocolor image during electrical stimulation at 2 Hz. The heart was loaded with the calcium-insensitive fluorophore calcein (5 µmol/l) and the fluorescent calcium indicator rhod-2 (10 µmol/l). Image was acquired at a scan rate of 1.33 ms/line during 2-photon excitation at 810 nm and emission was simultaneously measured in the green (500-550 nm) and red (560-650 nm) range. The image is an overlay of calcein and rhod-2 fluorescence signals, where calcein is shown in green and rhod-2 as red. Yellow indicates overlap between calcein and rhod-2 fluorescence. Electrical stimuli were continuously delivered to the right ventricular epicardium, resulting in cyclic increases in rhod-2 fluorescence along the length of the horizontal scan lines due to depolarization-induced rise in [Ca2+]i. White scale bar = 20 µm. Scaling was the same in x and y direction. B: pseudocolored line-scan image along the green line in A. This single line was scanned at a rate of 31.25 Hz. Separate line-scan images were constructed for the short- and long- wavelength emission range and subsequently combined to obtain the overlay image as shown. Red arrow demarks increase in stimulation rate from 1 to 2 Hz. Horizontal scale bar = 20 µm. Black downward arrow, 2 s. C: time courses of spatially averaged rhod-2 (red lines) and calcein (green lines) fluorescence from the 2 cells in B. Tracings were obtained by averaging pixel intensities in the horizontal direction and then plotting them as a function of time. There were stimulus-induced increases in rhod-2, but not calcein, fluorescence.



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  		Fig. 4.   Imaging of single myocyte calcium transients in the anterior left ventricle of a rhod-2-loaded heart at increasing tissue depth. A: time courses of spatially averaged, normalized rhod-2 fluorescence ratio (F/Fo) during electrical stimulation at either 1 or 2 Hz. Numbers indicate distance along the z-axis (in µm) starting from the epicardial surface. B: superimposed tracings of depolarization-induced, normalized changes in rhod-2 fluorescence at increasing depth as a function of time. Delta F were determined as described in Fig. 1F. Note the similarity in the time course of fluorescence changes at different depths.

Although the rhod-2 signal does not exhibit a specific pattern throughout the cardiomyocyte cytosol, there is a relatively heterogeneous tissue distribution of rhod-2 fluorescence in the heart. Strong fluorescent signals are present in small oval and elongated structures (see arrows in Figs. 1A and 5, A and B). The anatomic features of these structures (namely, the overall dimensions and their propensity to protrude into the capillary lumen) suggest that they correspond to endothelial cell nuclei (21). Epifluorescence microscopy of a 10-µm section from a heart loaded with rhod-2 and the nuclear counterstain Hoechst 34580 (Molecular Probes; Fig. 5, C and D) confirms that all areas of prominent red rhod-2 fluorescence also exhibit strong blue Hoechst fluorescence supporting the notion that they correspond to nuclei.


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  		Fig. 5.   Optical resolution of 2-photon molecular excitation (TPME) microscopy in mouse heart tissue. A and B: optical sectioning of left ventricular myocardium of a mouse heart loaded with rhod-2 using TPME fluorescence microscopy. Images were taken at intervals of 0.5 µm along the z-axis, starting at the epicardial surface in A and starting at 50 µm below the epicardial surface in B. Therefore, the lower left images in A and B are 10 and 60 µm into the ventricular wall, respectively. Rhod-2 fluorescence appears much more prominent in elongated and oval structures (white arrows) than in the surrounding cardiomyocytes. Red crosses mark positions of the cross-sectional slices through the image stacks in the x/z and y/z planes, which are shown in A and B, bottom. Images were collected at a pixel resolution of 430 × 430 × 500 nm in x, y, and z, respectively. Gain setting of the photomultiplier tubes was adjusted so that the F of the endothelial cells was below saturation and the background was close to zero. No background subtraction or image enhancement was used. Scaling was the same in the x and y direction. C and D: epifluorescence images of a 10-µm section from a heart loaded with rhod-2 and the nuclear counterstain Hoechst 34580. Note that all intensely red fluorescent structures in C (rhod-2 fluorescence) also exhibit prominent Hoechst fluorescence (blue; D). E and F: TPME images of a heart loaded with the fluorescent indicator di-4-ANEPPS taken at 10 and 80 µm, respectively, below the epicardial surface. Images were inverted for clarity. F was encoded in levels of gray. Regularly spaced T-tubule invaginations are visible. Black horizontal scale bar = 10 µm. Scaling was the same in the x and y direction. G: plot of di-4-ANEPPS Fs along the green and red lines in E and F.

We took advantage of the prominent rhod-2 staining in capillary endothelial cell nuclei to further evaluate the optical sectioning capability of TPME. Figure 5, A and B, shows full-frame images taken at 0.5-µm intervals along the z-axis through the left ventricle of a rhod-2-loaded heart; the arrows demark the intensely fluorescent nuclei of endothelial cells. Images in Fig. 5A are at a depth of 0 to 10 µm from the epicardial surface, whereas images in Fig. 5B are at a depth of 50 to 60 µm (corresponding to 4 layers of cardiomyocytes). As demonstrated in the cross-sectional slices through the image stacks in the x/z and y/z planes, the endothelial cell nuclear signals appear bright only over a distance of 1.5 to 2.0 µm along the optical axis and disappear within 3.5 to 4.0 µm (as the plane of focus is moved deeper into the specimen). This finding is consistent with the ellipsoid shape and dimensions of endothelial cell nuclei [average short diameters of ~2 µm (21, 42, 47, 48)] and the x/z-point spread function of the TPME laser scanning microscope (13, 46, 53). The x/z series also demonstrate that the axial resolution of the imaging system does not diminish noticeably over a distance from 0 to 60 µm along the optical axis.

To estimate the lateral resolution of our imaging system at increasing tissue depth, a mouse heart was loaded with the plasma membrane-selective indicator di-4-ANEPPS. TPME images obtained at 10 and 80 µm, respectively, below the epicardial surface are shown in Fig. 5, E and F. Di-4-ANEPPS produced periodic lines of fluorescence, corresponding to T-tubule invaginations (22). The distances between the peak fluorescence intensities of these lines (Fig. 5G) were virtually identical at tissue depths of 10 and 80 µm (2.32 ± 0.062 and 2.34 ± 0.050 µm, respectively; P > 0.05, t-test), indicating that the lateral resolution of the imaging system is 2.3 µm or less. As is the case for the axial resolution, the x/y resolution does not appear to diminish noticeably with increasing depth.

Although the point spread functions of our imaging system have not been determined using subresolution fluorescent beads (see Ref. 46), the x/z and y/z cross-sectional images clearly demonstrate that the thickness of the focal slice from which fluorescence signals are obtained is considerably less than the average depth, width, or length of adult murine ventricular cardiomyocytes [~13, ~32, and ~140 µm, respectively (see Ref. 44)]. This observation, in combination with the estimates of the lateral resolution, suggests that the system allows selective visualization of [Ca2+]i in a volume significantly less than that of a single cardiomyocyte within the intact heart without being affected by signals from neighboring cells.

Imaging of single myocyte [Ca2+]i transients in the intact mouse heart with fura-2. A similar series of analyses were performed using fura-2 to image [Ca+2]i in intact retrograde perfused hearts. Figure 6 illustrates electrically evoked changes in fura-2 fluorescence during TPME illumination at 800 nm in the presence of cytochalasin D and acetylcholine. As can be seen in the full-frame image (Fig. 6A), the resting fura-2 fluorescence is fairly homogeneously distributed within the cardiomyocyte cytosol, but it is relatively more intense in cardiomyocyte nuclei (Fig. 6A, arrowheads). All cardiomyocytes respond to pacing with a synchronous reduction of cytosolic fluorescence, corresponding to increases in cytosolic concentration of free calcium due to depolarization-induced activation of SR calcium release. The subsequent rise in fura-2 fluorescence during the decline in [Ca2+]i also occurs quite synchronously along the scan lines.


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  		Fig. 6.   Stimulation-evoked single myocyte [Ca2+]i transients in the intact left ventricle of a mouse heart loaded with fura-2. A: full-frame false color image during electrical stimulation at 1 Hz. Image was acquired at a scan rate of 1.33 ms/line during 2-photon excitation at 800 nm and emission was recorded in the 500-550 nm range. Intensity of fura-2 fluorescence recorded in the 500-550 nm range was encoded off-line in shades of green. Red arrowheads demark cardiomyocyte nuclei. Simultaneous decreases in fura-2 fluorescence along the length of the horizontal scan line are apparent in the middle of the image, corresponding to a depolarization that occurred when the scan was approximately halfway across the cells in the field of vision. White scale bar = 20 µm. Scaling was the same in the x and y direction. B: false color line-scan image along the red line in A. Scan speed, 32 ms/line. *Beginning and end of electrical stimulation at 2 Hz. Scale bars = 10 µm horizontally, 600 ms vertically. Black arrow indicates direction of vertical time axis. C: time courses of spatially averaged, normalized fluorescence ratio (F/Fo) from the regions indicated in B. Fo is the background intensity of fura-2 obtained by measuring F before the stimulus. Regions were 34, 29, and 18 µm wide, each one representing 1 of 3 juxtaposed cells along the scan line. The numbers in C correspond to the numbers of the cells in B. D: superimposed tracings of normalized changes in [Ca2+]i as a function of time from the 3 regions indicated in B. Delta F were determined as described in Fig. 1F. Note the similarity in the time course of fluorescence changes.

Synchronization of electrically evoked calcium transients among neighboring cardiomyocytes is also apparent in images obtained in line-scan mode during pacing at 1 and 2 Hz (Fig. 6B). Upon stimulation, fura-2 fluorescence decreases simultaneously within the first pixel in all cardiomyocytes along the horizontal scan line and then slowly returns to its prestimulus value. The time courses of spatially averaged fura-2 fluorescence (Fig. 6, C and D) confirm, in quantitative terms, the uniformity of changes in the fluorescence signals seen in juxtaposed cardiomyocytes. As was the case for rhod-2-loaded hearts, electrically evoked fura-2 fluorescence transients were readily recorded at depths up to 70 µm from the epicardial surface. The relative intensity (Fig. 7A) and normalized kinetics (Fig. 7B) of fura-2 transients recorded at different tissue depths are almost identical.


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  		Fig. 7.   Imaging of single myocyte calcium transients in the anterior left ventricle of a fura-2-loaded heart at increasing tissue depth. A: time courses of spatially averaged, normalized fura-2 fluorescence ratio (F/Fo) during electrical stimulation at either 1 or 2 Hz. Numbers indicate distance along the z-axis (in µm) starting from the epicardial surface. B: superimposed tracings of depolarization-induced, normalized changes in fura-2 fluorescence at increasing depth as a function of time. Delta F were determined as described in Fig. 1F. Note the similarity in the time course of fluorescence changes at different depths.

The emission spectra of calcium-free and calcium-bound fura-2 are almost superimposable with maxima at 512 and 505 nm, respectively (38). Although the two-photon cross section of calcium-bound fura-2 at an excitation wavelength of 800 nm is reportedly two orders of magnitude smaller than that of calcium-free fura-2 (56), it remains a possibility that the fluorescence of both calcium-free and calcium-bound fura-2 contributes to the signal in the 500-550 emission range (with the decrease in fluorescence predominating). However, decreasing the two-photon excitation wavelength from 800 to 740 nm has no significant effect on the ratio F/Fo or its kinetics (Fig. 8), suggesting that calcium-bound fura-2 is not excited under the conditions employed.


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  		Fig. 8.   Fura-2 transients during 2-photon excitation at 2 different wavelengths. Time course of changes in F/Fo (means ± SE; n = 5 cells from 3 hearts/group) in fura-2-loaded hearts (1 Hz) during 2-photon illumination at wavelengths of 740 and 800 nm. At some time points, symbols are larger than SE bars. F/Fo curves were almost superimposable.

Comparison of electrically evoked fura-2 and rhod-2 fluorescence transients. Figures 1 and 6 illustrate marked differences in the apparent rate of decline of intracellular free calcium as estimated by the relative changes in rhod-2 vs. fura-2 fluorescence. This difference is quantitated in Table 1, which summarizes the properties of electrically evoked rhod-2 and fura-2 transients. With either fluorophore, there is a slight but significant change in the prestimulus fluorescence intensity with an increase in stimulation rates (see Fo2Hz/Fo1Hz values, Table 1). In addition, doubling the stimulation rate causes a statistically significant shortening of the early and late relaxation phase of the F/Fo transients, as documented by the decrease in average t90-50% and t50-10% values. The time course of [Ca2+]i decline was apparently slower in fura-2-loaded cardiomyocytes than in rhod-2-loaded cardiomyocytes. Collectively, these data indicate that while both fluorophores can faithfully track increases in [Ca2+]i changes that occur with depolarization, there is some discrepancy in their ability to monitor the decreases in [Ca2+]i that occur during repolarization. It is also important to note that F/Fo values between 1.5 and 2.5 in rhod-2-loaded hearts and between 0.5 and 0.3 in fura-2-loaded hearts could occasionally be measured during spontaneous calcium transients. This latter observation indicates that changes in cytoplasmic Ca2+ occurring under the conditions here are within the effective range of rhod-2 for measuring Ca2+ levels and, furthermore, that changes in the fluorescence of Ca2+-free fura-2 were above background levels during pacing at 1 and 2 Hz.

The magnitude and kinetics of the changes in intracellular calcium depend on the calcium-buffering properties of the cytoplasm (18, 36). In addition to endogenous buffers, fluorescent calcium indicators have been reported to have effects on calcium buffering (5, 18). To directly test the hypothesis that the slower decay of the evoked calcium transient in fura-2-loaded cardiomyocytes compared with rhod-2-loaded cardiomyocytes is due to the increased buffering capacity of fura-2, the effects of extra buffering by the nonfluorescent fura-2 analog BAPTA on the time course of the rhod-2 transient were measured. As expected, incorporation of BAPTA (5 µmol/l) slows the rate of decay of the calcium transient (Fig. 9, A and B). Spatially averaged calcium transients in Fig. 9A also demonstrate an increase in prestimulus calcium with an increase in stimulation frequency due to incomplete recovery of the calcium transient during the shorter interpulse interval. Superimposition of normalized fluorescent transients (Fig. 9B) reveals that incorporation of BAPTA causes the kinetics of rhod-2 decay to become virtually identical to those obtained with fura-2. Table 1 summarizes the effects of adding BAPTA on the peak and decay rate of the stimulated rhod-2 transient. Extra buffering by BAPTA significantly decreases both the peak and decay of the rhod-2 transient. The increase in prestimulus rhod-2 fluorescence associated with the increase in stimulation rate from 1 to 2 Hz is also more pronounced in the presence of BAPTA. BAPTA prolongs the early relaxation to a similar degree as fura-2, but it slows the late phase of decay to a significantly higher extent.


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  		Fig. 9.   Effect of extra buffering of intracellular calcium by BAPTA on electrically evoked rhod-2 transients. A: time courses of spatially averaged, normalized rhod-2 fluorescence, F/Fo, from 3 juxtaposed cardiomyocytes during electrical stimulation at either 1 or 2 Hz. The heart was loaded simultaneously with the calcium indicator rhod-2 (10 µmol/l) and the nonfluorescent calcium buffer BAPTA (5 µmol/l), and F was measured in the 560-650 nm range. B: superimposed tracings of changes in single cardiomyocyte [Ca2+]i as a function of time from hearts loaded with fura-2 or rhod-2 in the absence and presence of BAPTA, respectively. Delta F were determined as described in Fig. 1F.

TPME imaging of abnormalities in intracellular calcium handling in intact mouse hearts. Abnormalities in intracellular calcium handling have long been implicated as the underlying mechanism in a number of pathological conditions that promote arrhythmia. For example, cellular calcium overload may initiate sporadic calcium waves, which in turn induce proarrhythmogenic afterdepolarizations. Occasionally, elevations and reductions, respectively, of rhod-2 and fura-2 fluorescence occurred between stimulated or spontaneous [Ca2+]i transients, consistent with the appearance of sporadic calcium waves. Figure 10A shows a rhod-2 fluorescent image from the left ventricular anterior wall at a depth of ~30 µm in a Langendorff-perfused heart. The calcium waves exhibited intracellular propagation, similar to those observed in single myocytes and multicellular cardiac preparations (10, 27, 32, 54). Waves were detectable when the sample was paced at 1 Hz (Fig. 10A, waves a and b), 2 Hz (Fig. 10A, wave c), and also during spontaneous depolarization (Fig. 10A, waves d-f). Occasionally, intracellular truncation of a calcium wave occurs (see, for example, wave f in Fig. 10A). High-frequency calcium waves were also observed. For example, waves initiating at a frequency of ~180/min (as calculated from the line-scan image) are depicted in Fig. 10B. These high-frequency waves show no propagation to adjacent cardiomyocytes. Similar high-frequency calcium waves have previously been reported in fluo-3-loaded, buffer-perfused rat hearts (27). Thus the TPME system is able to detect temporal heterogeneity of [Ca2+]i signaling with subcellular resolution in the stimulated and spontaneously depolarizing intact heart.


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  		Fig. 10.   Spontaneous calcium waves in the Langendorff-perfused mouse heart. A: line-scan image along 2 adjacent myocytes in a heart loaded with rhod-2 during incremental stimulation from 1 to 2 Hz. Image is shown by pseudocolors corresponding to the rhod-2 F, with black being the lowest and red being the highest intensity. Collisions of spontaneously developing calcium waves with calcium transients evoked by electrical stimulation caused annihilation of the waves (waves a, b, and c), whereas waves d, e, and f terminate spontaneously or disappear from the plane of focus. None of the waves a through f spreads to the neighboring cell. Average propagation velocity of the calcium waves was ~120 µm/s. #Spontaneous depolarization-induced calcium transients. Scale bars = 10 µm horizontally, 500 ms vertically. B: high-frequency Ca2+ waves during electrical stimulation (1 Hz) in 2 neighboring cardiomyocytes in a fura-2-loaded heart. Image is shown by pseudocolors corresponding to the fura-2 F. Frequency of the waves is 180 min-1. None of the Ca2+ waves in cell 1 transmits to neighboring cell 2. Scale bars = 5 µm horizontally, 500 ms vertically.

Abnormalities in intracellular calcium handling may also give rise to mechanical alternans, which are a predictor of fibrillation (35). We previously generated a transgenic mouse model that has constitutively elevated levels of TGF-beta 1 activity in the adult heart. These animals develop progressive atrial fibrosis, but the ventricles are not adversely affected by transgene expression (Ref. 34; Fig. 11, A and B). Intracardiac electrophysiological studies revealed that these animals exhibit increased susceptibility to electrically induced atrial fibrillation compared with their nontransgenic controls (43), leading to the suggestion that atrial fibrosis results in the formation of an arrhythmogenic substrate. We therefore studied the atria from intact hearts from these mice with TPME in an effort to determine if cellular heterogeneity in calcium handling might also be present. In the left atrium of the TGF-beta 1-expressing mouse, there was a marked alternans of electrically induced [Ca2+]i transient amplitude in one cardiomyocyte (cell 2) but not in its immediate neighbors (Fig. 11D). Thus the system described here is also capable of detecting spatial heterogeneity of [Ca2+]i signaling within the intact heart.


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  		Fig. 11.   [Ca2+]i transient alternans in a single atrial cardiomyocyte in the heart of a TGF-beta 1 transgenic mouse. A and B: histological sections of left atria from a 4-mo-old TGF-beta 1 transgenic mouse (B) and its wild-type littermate. Sections were stained with Sirius red and fast green. Note the abundant collagen deposition (red signal) throughout the transgenic atrial myocardium. C and D: time courses of spatially averaged, normalized rhod-2 fluorescence, F/Fo, from 4 juxtaposed cardiomyocytes each in the left atrium of a 4-mo-old transgenic mouse with cardiac-specific expression of a constitutively active TGF-beta 1 (D) and in the left atrium of a nontransgenic littermate. Rhod-2 transients were recorded during right atrial stimulation at either 1 or 2 Hz. Note the presence of a marked alternans of calcium transient amplitudes (red arrows) in 1 cardiomyocyte in the transgenic atrium but not in its immediate neighbors.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

In this study, we describe a technique using TPME in combination with laser scanning microscopy to measure [Ca2+]i-dependent changes in the fluorescence intensity of calcium indicators at the single cardiomyocyte level in buffer-perfused mouse heart preparations. The technique permits quantitative assessment of stimulation-evoked calcium transients at intramyocardial depths <= 100 µm from the epicardial surface. This imaging system clearly demonstrates that calcium-induced calcium release activated by depolarization and the removal of calcium from the cytosol are highly synchronized among neighboring cardiomyocytes under our experimental conditions. The system is also sensitive enough to detect the presence of calcium waves and [Ca2+]i alternans in the paced heart. The axial and lateral resolutions do not diminish appreciably with increasing depth into the tissue for TPME illumination (13, 57), at least over a distance of 60 µm from the epicardial surface. We further demonstrate that the widely used excitation-contraction uncoupler BDM cannot be used for TPME imaging of stimulated calcium transients due to its adverse effects on cardiac excitability. On the other hand, cytochalasin D not only effectively uncouples contraction from excitation, but it also retains physiological phenomena of cardiac calcium signaling, such as rate-dependent acceleration of [Ca2+]i decline.

Although we have not unequivocally determined the subcellular localization of rhod-2 and fura-2 in our study, examination of the full-frame images strongly suggests that both indicators accumulated in the nuclei and cytosol of cardiomyocytes. Relatively homogeneous fluorescence patterns are present in the cytosol of quiescent, rhod-2- or fura-2-loaded cardiomyocytes [as opposed to the more punctuate appearance typically observed with mitochondrial fluorescent labels (9, 16, 33, 51)]. This is in agreement with the results of previous studies that employed dye loading protocols similar to those used here. For example, Del Nido and colleagues (16) showed that rhod-2 was located in the cytosol with no evidence of mitochondrial deposition in isolated guinea pig ventricular myocytes loaded with rhod-2/AM. Similarly, MacGowan et al. (31) recently confirmed the absence of mitochondrial rhod-2 accumulation in murine ventricular myocytes, using both electron and fluorescence confocal microscopy, and Zoghbi et al. (58) also found no evidence for mitochondrial rhod-2 uptake in isolated rat ventricular myocytes. Other groups (24, 33, 50, 51) found rhod-2 accumulation predominantly in the mitochondria of cardiomyocytes. This apparent discrepancy is most likely due to the use of very different loading protocols in these studies. The observation that rhod-2 fluorescence transients evoked by electrical stimulation completely abolished propagating elevations in indicator fluorescence (previously dubbed calcium waves) is consistent with the notion of a preferential cytosolic (extramitochondrial) localization of rhod-2. Calcium waves have been shown to result from propagating activation of neighboring clusters of ryanodine receptors in the SR membrane. Annihilation of calcium waves by colliding intracellular transients is thought to be due to ryanodine receptor refractoriness in the wake of a wave of calcium release (6, 10). Such interaction could not occur if the electrically evoked increase in indicator fluorescence were confined to the mitochondria or other organelles.

The data obtained with TPME are similar to those of other systems. Du et al. (19) previously measured calcium-dependent changes in rhod-2 fluorescence of whole mouse heart. Under their experimental conditions (37°C, stimulation at 8 Hz, [Ca2+]o 2.5 mmol/l), systolic F/Fo peaked at ~1.3 (see Fig. 6b in Ref. 19). This value compares surprisingly well with our measurements at the single cardiomyocyte level within intact tissue, despite the marked differences in experimental conditions (i.e., perfusate temperature and calcium concentration, stimulation frequency, presence of acetylcholine and cytochalasin D). Similarly, Ito et al. (25) studied [Ca2+]i regulation at 25°C and 1.5 mmol/l perfusate calcium in mouse myocytes loaded with the calcium indicator fluo-3/AM. Raising the stimulation frequency from 1 to 2 Hz caused increases in diastolic and peak systolic [Ca2+]i and marked acceleration of the decline in [Ca2+]i. This behavior strikingly resembles that of fura-2 and rhod-2 fluorescence transients in this study, indicating that the rate-dependent dynamics of intracellular calcium regulation in situ replicate those in isolated myocytes in vitro.

It is of interest to note that the fura-2 fluorescence signal during stimulus-evoked transients returned to its prestimulus value much more slowly than the rhod-2 signal. Fluorescent indicators that are used to measure [Ca2+]i have been reported to have effects on cytoplasmic calcium buffering, resulting in a reduction of both the peak and rate of decay of the systolic calcium transients (18). Incorporation of the nonfluorescent calcium buffer BAPTA, which is structurally related to fura-2 and has a similar Ca2+ affinity, decreased peak amplitude and decay rate of stimulated rhod-2 transients in our study. Extra buffering by the calcium chelator nitr-5, which has similarly high affinity for calcium and similar kinetics of binding and release of calcium as BAPTA, has previously been shown to reduce both peak and decay rate of systolic calcium transients in single rat ventricular cardiomyocytes (18). In the latter study, the effects of nitr-5 on amplitude and kinetics could be attributed quantitatively to increased calcium buffering. Thus our findings strongly support the notion that, in the range of concentrations used in our experiments, fura-2 buffers cytosolic calcium in cardiomyocytes, resulting in smaller amplitude and slowed relaxation of the calcium transient. Consistent with these interpretations is the previous observation by Backx and ter Keurs (2) that the decay rate of the Ca2+ transient is inversely related to the intracellular fura-2 concentration in iontophoretically loaded rat papillary muscles. Collectively, these studies emphasize the importance of knowing the possible effects that fluorescent calcium indicators have on the time course and magnitude of the physiological events under study, if quantitative interpretation of the signal is to be obtained.

The use of cytochalasin D at relatively high concentrations to effectively uncouple contraction from excitation constitutes a potential limitation for the approach described here. Although cytochalasin D at a concentration of 40 µmol/l has previously been found to have no effects on the kinetics of calcium transients in isolated rat ventricular myocytes, a slight increase in the peak amplitude was noted (23). The effects of cytochalasin D on [Ca2+]i transients in mouse ventricular myocytes are currently unknown. A previous study demonstrated that cytochalasin D markedly prolongs action potential duration in murine ventricular myocardium in a concentration-dependent manner (3). Changes in membrane potential would affect the activities of voltage-dependent calcium conductances, leading to changes in the amplitude and/or kinetics of [Ca2+]i transients.

Use of the alternative excitation-contraction decoupler BDM is not feasible due to its adverse effects on cardiac excitability at concentrations that are necessary to eliminate motion (Fig. 2). Because other pharmacological agents [such as L-type calcium channel blockers (45) or calcium chelators (41)] eliminate or significantly reduce calcium transients, they were not suitable for the analyses performed here. Similarly, mechanical immobilization significantly alters epicardial activation patterns during voltage-sensitive dye mapping of Langendorff-perfused rat hearts (37) and is of limited utility. Thus, although cytochalasin D may exert as yet undefined minor effects on the time course and magnitude of stimulated calcium transients, our observation that the kinetics of calcium transients and their frequency dependence in cytochalasin D-treated cardiomyocytes are qualitatively very similar to those in isolated, untreated myocytes (1, 25) suggests that the use of this fungal metabolite does not preclude the study of physiological phenomena.

Acetylcholine may also affect the properties of intracellular calcium transients by means of direct interaction with calcium channels/transporters and/or via modulation of the action potential, which in turn controls the activity of voltage-dependent calcium conductances (28). However, with the availability of microscopes with higher scan speeds, it will likely become possible to image [Ca2+]i transients in the intact mouse heart at its intrinsic rate (and at more physiological temperatures), rendering the use of negative chronotropic substances unnecessary. Finally, we did not determine the two-photon excitation spectra or the two-photon cross-sectional areas of rhod-2 and fura-2. It is possible that wavelengths below or beyond those provided by our laser source may enhance imaging capabilities by way of improving tissue penetration depth or increasing the likelihood of two-photon excitations.

The system described here permits imaging of stimulation-evoked calcium transients with subcellular resolution in individual cardiomyocytes at tissue depths that have previously been inaccessible to single-photon laser-scanning confocal microscopy. As demonstrated above, the approach should be of general utility to monitor the consequences of genetic and/or functional heterogeneity between neighboring cardiomyocytes. For example, spontaneous increases in intracellular calcium concentration in cardiomyocytes are thought to trigger afterdepolarizations, which then transmit to neighbor cells, ultimately launching arrhythmias. Afterdepolarizations have long been hypothesized to trigger arrhythmias in the acutely ischemic myocardium and in hearts of long QT patients (14). Spatially heterogeneous [Ca2+]i transient alternans has previously been demonstrated in ischemic hearts (55), where they are of apparent importance for the development of arrhythmias. The technique presented here provides the unique opportunity to define the cell-cell interactions that promote intercellular propagation of [Ca2+]i-dependent afterdepolarizations in the whole heart under these pathological conditions. The use of TPME, in conjunction with the ability to genetically modify mice at will through the use of transgenesis or gene targeting, constitutes a very powerful experimental system. This technique is also particularly well suited to follow the functional fate of donor cells following direct intracardiac transplantation (40) or following homing to sites of injury (26), provided that the donor and host cells can be discriminated on the basis of fluorescent properties. The ability to monitor functional parameters at the individual cell level (as opposed to global heart function) may also permit a better assessment of the effects of donor cells following transplantation into injured hearts (that is, discriminating between a nonspecific effect on postinjury remodeling vs. a direct contribution to a functional syncytium).


    ACKNOWLEDGEMENTS

We thank J. D. Dowell and Drs. K. Pasumarthi, M. Soonpaa, H. O. Nakajima, and H. Nakajima for helpful comments on the manuscript.


    FOOTNOTES

This study was supported by National Institutes of Health grants (to L. J. Field).

Address for reprint requests and other correspondence: M. Rubart, Wells Center for Pediatric Research, Riley Hospital, 702 Barnhill Drive, Indianapolis, IN 46202 (E-mail: mrubartv{at}iupui.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

First published February 12, 2003;10.1152/ajpcell.00469.2002

Received 3 October 2002; accepted in final form 10 February 2003.


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DISCUSSION
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