Proc. Natl. Acad. Sci. USAVol. 93, pp. 5413-5418, May 1996Physiology

Intrasarcomere [Ca2+J gradients in ventricular myocytes revealedby high speed digital imaging microscopyGERRIT ISENBERG*, ELAINE F. ETrERt, MARIA-FLORA WENDT-GALLITELLI*, ALFRED SCHIEFER*,WALTER A. CARRINGTONt, RICHARD A. TUFTt, AND FREDRIC S. FAYt*Martin-Luther-Universitat, Halle-Wittenberg, Medizinische Fakultat, Julius-Berstein-Institute fur Physiologie, Magdeburger Str. 6, 06097 Halle/S., Germany; andtDepartment of Physiology, Biomedical Imaging Group, University of Massachusetts Medical School, Biotech Two, 373 Plantation Street, Worcester, MA 01605

Communicated by Erwin Neher, Max-Planck-Institut fuir Biophysikalische Chemie, Gottingen, Germany, December 11, 1995 (received for reviewAugust 28, 1995)

ABSTRACT Cardiac muscle contraction is triggered by asmall and brief Ca2+ entry across the t-tubular membranes,which is believed to be locally amplified by release of Ca2+from the adjacent junctional sarcoplasmic reticulum (SR). AsCa2+ diffusion is thought to be markedly attenuated in cells,it has been predicted that significant intrasarcomeric [Ca2+]gradients should exist during activation. To directly test forthis, we measured [Ca2+] distribution in single cardiac myo-cytes using fluorescent [Ca2+] indicators and high speed,three-dimensional digital imaging microscopy and image de-convolution techniques. Steep cytosolic [Ca2+] gradients fromthe t-tubule region to the center of the sarcomere developedduring the first 15 ms of systole. The steepness of these [Ca2+]gradients varied with treatments that altered Ca2+ releasefrom internal stores. Electron probe microanalysis revealed aloss of Ca2+ from the junctional SR and an accumulation,principally in the A-band during activation. We propose thatthe prolonged existence of [Ca2+] gradients within the sar-comere reflects the relatively long period ofCa2+ release fromthe SR, the localization of Ca2+ binding sites and Ca2+ sinksremote from sites of release, and diffusion limitations withinthe sarcomere. The large [Ca2+I transient near the t-tubular/junctional SR membranes is postulated to explain numerousfeatures ofexcitation-contraction coupling in cardiac muscle.

In cardiac muscle, upon stimulation, a small Ca2+ influxthrough voltage-gated Ca2+ channels in the t-tubular mem-branes (1) is greatly amplified by a Ca2+-induced release ofCa2+ through ryanodine receptor/Ca2+ channels on the ad-jacent junctional sarcoplasmic reticulum (2). Because Ca2+diffusion inside cells is greatly attenuated by an abundance ofCa2+-buffering proteins (3), significant intrasarcomeric [Ca2+]gradients have been predicted to exist during Ca2+ release incardiac myocytes (4). Recent studies using confocal microfluo-rimetry have revealed the existence, following activation, of asmall and transient intrasarcomeric [Ca2+] gradient in frogskeletal muscle emanating from regions of the sarcomerecontaining junctional sarcoplasmic reticulum (SR) (5). Theseobservations support earlier autoradiographic studies (6) thatsuggested that these elements were responsible for Ca2+release following membrane depolarization. It remains to beseen whether similar gradients of [Ca2+] exist during activationof cardiac muscle where release of Ca2+ from the SR ismediated by a somewhat different and slower mechanism (7)involving a different class of ryanodine receptors (8) coupledto membrane depolarization by a local rise in [Ca2+] (9) ratherthan a mechanical mechanism as is thought to operate inskeletal muscle (10). Recent studies using confocal laserphotometry have revealed the existence of brief and transientmicroscopic domains of high [Ca2+] in cardiac myocytes, which

have been interpreted as the elementary events of SR Ca2+release (11). In order to visualize these events and theirrelationship to the opening of single voltage-controlled Ca2+channels, these studies were performed under conditionswhere full excitation-contraction coupling was suppressedeither by use of Ca2+ channel blockers or by using very briefdepolarizations (12). The local [Ca2+] changes underlyingexcitation-contraction coupling in heart muscle under morephysiological conditions have, however, not been examined.Nor is there information about how possible patterns of [Ca2+]changes following depolarization are related to the organiza-tion of elements underlying excitation-contraction coupling incardiac muscle. The current studies were thus carried out toprobe for the existence of [Ca2+] gradients and their relation-ship to structures believed to underlie excitation-contractioncoupling during rhythmic contractions of heart cells underphysiological conditions.

METHODSCell Isolation, Electrophysiology, and Dye Loading. Myo-

cytes were isolated from the ventricles of 400-g guinea pigs asdescribed previously (13). Fluorescent Ca2+ indicators wereloaded into single voltage-clamped guinea pig cardiac myo-cytes through a patch pipette. Cells in recording chamber werecontinuously superfused with prewarmed (37°C) physiologicalsalt solution composed of 150mM NaCl, 2 mM CaCl2, 5.4mMKCI, 1.2 mM MgCl2, 20 mM glucose, 10 mM Hepes/NaOH(pH 7.4). Cells were voltage clamped with patch electrodes of2-3 Mfl resistance. Electrodes were filled with 140 mM KCl,10mM NaCl, 10mM Hepes/KOH (pH 7.2), 0.1 mM K5Fura-2or 0.1 mM calcium green-2. Cells were rhythmically stimulatedby single or paired voltage clamp pulses repeated every second.Starting from a holding potential of -45 mV (to inactivate thefast sodium current), 180-ms steps to +10 mV were madefollowed by 820-ms diastolic repolarization. For the pairedpulse protocol, a second 180-ms pulse to +50 mV followed thefirst one after a 20-ms repolarization to -45 mV; this protocolis thought to optimally load the sarcoplasmic reticulum withCa2+ via Ca2+ influx through the Na+-Ca2+ exchanger (14).Measurements of [Ca2l] by Fluorescence Imaging. In order

to image the [Ca2+] distribution during systole, images of thefluorescence of either calcium green or Fura-2 were collectedjust prior to depolarization and then at times (4-6, 8-10, and12-14 ms) after the start of a pulse to + 10 mV, correspondingto the plateau of the cardiac action potential. Following a trainof 10 stimulating single or paired pulses (described previously),images were acquired at the start of depolarizing pulses to + 10mV that repeated every second (five repeats). The focal planewas changed between voltage pulses, each second, so that timeseries of images were acquired at five focal planes through the

Abbreviations: SR, sarcoplasmic reticulum; 3-D, three-dimensional;TR-WGA, Texas Red-labeled wheat germ agglutinin.

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The publication costs of this article were defrayed in part by page chargepayment. This article must therefore be hereby marked "advertisement" inaccordance with 18 U.S.C. §1734 solely to indicate this fact.

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cell. The entire protocol was repeated at a second wavelengthwhen Fura-2 was used to measure [Ca21]. These images werereorganized, and each set of five optical sections correspond-ing to a time relative to depolarization was processed using aconstrained deconvolution algorithm to reassign light to itspoint of origin in 3-D. This process provides a more accurateimage of fluorescence at all focal planes and in turn allows formore accurate estimates of [Ca2+] (15).

[Ca2+] was calculated point-by-point from the processedimages of Fura-2 fluorescence using the following expressions:

2+aFua- =KDI[R/Rdia - (Rmin/Rmax)C][Ca2]Fura-2 = C-R/Rdia [1]whereR = F351/F380 and Rdia is the fluorescence ratio just priorto depolarization, and

KD!3 + [Ca2 ]diaC [Ca2+Idia + KDI3(Rmin/Rmax) [2]

[Ca2+]dia is assumed to be uniformly 120 nM in a cell at rest,150 nM in a cell at diastole following 10 single stimulatingpulses, and 170 nM in a cell at diastole following stimulationwith 10 double pulses (13). Rmin and Rma are globally mea-sured values of R in the absence of Ca2+ and in saturating[Ca2+], respectively; 13 = F380 in the absence of Ca2+/F380 insaturating [Ca2+], and KD was taken as 200 nM (16). Thisexpression was utilized to calculate [Ca2+] from the 351- and380-nm excitation images in order to correct for small differ-ences in the spatial patterns of illumination at these twowavelengths. These differences arose due to minor differencesin the interference of the two separated coherent light beamsthat had slightly different paths through the microscope optics(Fig. 1). This equation is simply derived from the standardratiometric equation (16) and makes use of the fact that whileRmjn andRm. will vary slightly due to differences in the patternof 351- and 380-nm excitation, the ratio ofRmin/Rm. and ,B areinherent properties of the dye, unaffected by spatial variationsin excitation at the two wavelengths. The only additionalassumption underlying this equation is that [Ca2+] prior tostimulation is uniform and its value is known (13). Thefluorescence at both wavelengths bleached to a variable extentduring an experiment, and because a fluorescent intermediatewith spectral properties similar to Ca2+-free Fura-2 is pro-duced during this process (17), an uncorrected downward biasin the calculated [Ca2+] was superimposed on the elevation of[Ca2+] during depolarization; similar effects were not apparentin the calcium green records. [Ca2+] was calculated point bypoint from the processed images of calcium green fluorescenceas follows:

[Ca2]ca-green-2KD[F(a[Ca2+Idia/KD + 1) -Fd(l + [Ca2+Idia/KD)IaFd(l + [Ca2Idia/KD)- F(a[Ca2dia/KD + 1) [3]

where Fd = fluorescence at diastole, a = F in saturating[Ca2+]/F in absence of Ca2+, and KD was taken to be 500 nM.This expression was derived using an approach similar to thatused for other nonratiometric dyes in cardiac cells (11). Allvalues reported are the mean ± SE.

Electron Probe Microanalysis. Electron probe microanaly-sis was used to determine the distribution of total calcium insingle cardiac muscle cells quick frozen in a defined physio-logical state as described previously (14). Briefly, when pairedpulses had potentiated the contraction to the maximum, singlecardiac muscle cells were shock frozen with propane (- 196°C).Shock freezing was timed at the end of diastole or 10-15 msafter the start of depolarization. The frozen cells were cut at

FIG. 1. Schematic illustration of the partial-confocal laser-illuminated microscope system used to collect 3-D images of Fura-2 orcalcium green-2 fluorescence. Upper left, membrane current andexposure times for image acquisition typically used to evaluate [Ca2+]transients following depolarization. Fluorescence was imaged at agiven focal plane for 3 ms at 10 ms before and at three time points (inthis example at 4, 8, and 12 ms) after depolarization to + 10 mV froma holding potential of -45 mV. The microscope system consists of awavelength tuneable argon-krypton laser (Li), which was used to excitecalcium green-2 (A = 488 nm) or Texas Red (A = 563 nm). Fura 2fluorescence was excited by 351- and 380-nm lines of an argonmultiline UV laser (L2), which are split and recombined by 45° dichroicbeamsplitters (DI) and (D2); the lines are further isolated by bandpassfilters (Fi) and (F2). Two 3-mm diameter laser shutters (S) controlexposure durations. Illumination beams are combined by dichroicbeamsplitter (D3) and reflected onto the cell by epi-illuminationdichroic D4 through a lOOX numerical aperture, 1.3 UV objective. Aslit aperture (SA) limits the illumination to a 3 i,m wide strip on thecell, providing a partial confocal effect, which reduces the fluorescencesignal from out of focus regions of the cell. The slit was removed forthe wide field images of TR-WGA (see Fig. 3). The cell fluorescenceis imaged by the objective through emission bandpass filter (F3). Amask (M) in the image plane of the objective limits the reimagedfluorescence to a 25 x 512 pixel region on the 512 x 512 pixel scientificgrade charge-coupled device imager. The masked region of thecharge-coupled device (MR) stores prior images acquired in the timeseries which have been moved under the mask by parallel chargetransport (PCT). A piezoelectric translator (P) allows rapid focusingof the objective to 5 focal planes (-1.5, -0.5, 0, 0.5, 1.5 ,um) within thecell. A 386 PC controls all imaging and patch-clamp protocols.

- 150°C into 80-nm thin sections. The cryosections werefreeze-dried under high vacuum (10-5 Torr), carbon-coated,and analyzed in a Philips CM-12 STEM electron microscope.The x-ray spectra were processed by subtracting continuumcomponents from the grid, the holder, and the microscope.The location of the peak centroid and the resolution of thedetector were continuously calibrated by a computer-fittingroutine. Standards of known concentrations of potassium andcalcium were used for calibration. With a 1000-s analysis timeper spectrum, the minimum detectable calcium concentrationwas 0.08 mM. Quantitative data was statistically evaluated witha SAS statistic package using analysis of variance (followed byTukey-Kramer test). Electron probe microanalysis measuresthe total calcium concentration; i.e., the sum of bound andionized calcium. Total [Ca] was measured in longitudinalcryosections (80 nm thick before freeze-drying, see ref. 14). Toresolve the spatial distribution of total [Ca2+], the narrow areaof analysis ("scanning area"), which was 20 nm wide along the

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fiber axis and 700 nm long perpendicular to the fiber axis, wasscanned from z-line to z-line as described previously (14).Sarcomeres were selected for analysis where the A-band wasdevoid of longitudinal elements of the SR asjudged by the tightparallel alignment of thick filaments and the absence of astrong phosphorus signal characteristic of membranes.Computer Simulations. [Ca2+] distribution in one dimen-

sion during the onset of systole was computer-simulated by themethod of Kargacin and Fay (18), approximating the contin-uous space with 30 elements each 30 nm in size. Ca2+ releaseat a rate of 7 x 10-24 mol s-1 nm-2 was assumed to take placecontinuously (19) for 15 ms at the origin only for the simula-tions where Ca2+ buffers were present. For the simulationwhere no Ca2+ buffers were present in the model, the Ca2+release rate was adjusted downward by 1000 times so that themean [Ca2+] in the sarcomere was in the same range as thatin the presence of troponin C and myosin. Diastolic [Ca2+] wasset to 170 nM. Values used in the simulations were Ca2+, DCa(diffusion coefficient) = 7 x 10-6 cm2 s'-; Fura-2 (19), Dfura-2= 1.57 x 10-7 cm2 S-1, KdCa (dissociation constant) = 0.14,tM, kon = 9 X 108M-1 s-1, concentration 100 ,uM distributeduniformly; troponin C (20), KdCa = 2.15 ,uM, k0n = 1.2 x108M-1 S-1, concentration 400 ,tM distributed uniformly;myosin (20), KdCa = 0.83 mM, kon = 6 x 107 S-1, concentration800 ,uM, distributed uniformly between 0.1 and 0.8 ,um fromorigin. While the DCa used in the simulation was that in freesolution, the movement of Ca2+ within the sarcomere wasconsiderably slowed by the binding of Ca2+ to proteins, whosediffusion coefficient is considerably less than that of Ca2+alone. A 3-D model was created from this one-dimensionalsimulation, forming a volume 10.8 ,um x 7.7 ,um x 8 ,um withconcentrations varying only in the x direction and with sar-comeres 1.8 ,um long. Fluorescence at 351 and 380 nm wascalculated from the simulated values for [Fura-2] and [Fura-2-Ca2+] and empirically determined values for the relativebrightness of Fura-2 and Fura-2-Ca2+. These fluorescentmodels were then blurred by a point spread function thatincorporated the effect of using partial confocal slit illumina-tion; the results were binned to 0.12 ,um x 0.12 ,um pixel size.The deconvolution was performed on each wavelength imagewith smoothing parameters, a = 0.005 and with 30 iterations,identical to what was used to process images obtained fromactual cells (15). Note that since this simulation models a

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nonequilibrium situation, calculating [Ca2] from the fluores-cence ratios by the method of Grynkiewicz et al. (16) under-estimates the gradient of [Ca2+] and gives a value for the[Ca2+] gradient that is 40% of the value of the model.

RESULTS AND DISCUSSIONImages of [Ca2>] distribution at times following a depolarizingstep in two cardiac myocytes are shown in Fig. 2. In onemyocyte, [Ca2+] was estimated from fluorescence of Fura-2 attwo wavelengths (panel C). In the other, fluorescence ofcalcium green-2 at a single wavelength was utilized (panelA).Patterns in the transient changes in [Ca2+] recorded usingthese two indicators are quite similar. At 4 ms, the [Ca2+]increase was relatively small. By 8 and 12 ms, [Ca2+] hadincreased considerably, and large [Ca2+] gradients within thecell were evident, which exhibited a periodicity of 1.5-2.0 ,umalong the long axis of the cell, similar to the 1.8-p.m sarcomerespacing of relaxed isolated ventricular cells. In four cells inwhich [Ca2+] was measured with Fura-2 using the double pulseprotocol, the average [Ca2+] 4-5 ms after start of depolariza-tion was 151 ± 7 nM. By 8-10 ms, average [Ca2+] was 205 ±20 nM. On average, measuring 30 local [Ca2+] gradients, peak[Ca2+] was 373 ± 25 nM and [Ca2+] in the valleys between thepeaks was 132 ± 13 nM. By 12-15 ms after start of depolar-ization, the peaks of [Ca2+] were close to 1 p.M (897 ± 123nM); the valleys were 236 ± 35 nM, and average [Ca2+] was 362± 55 nM. Similar results were obtained from the analysis ofimages of calcium green-2 fluorescence (Fig. 2B). Mean valuesfor 9 cells: (spatially averaged [Ca2+]) 174 ± 5 nM (at 4-5 ms),273 ± 29 nM (8-12 ms), and 354 + 44 nM (12-15 ms). Periodic[Ca2+] gradients (peak/valley), mean values in nM: (407 ±45)/(228 + 24) (8-10 ms), and (606 ± 99)/(345 ± 51) (12-15ms). As can be seen from the contour presentation of the data(Fig. 2 E and F) the magnitude of the peaks of [Ca2+] variedwithin the cell; in some regions large peak [Ca2+] increasesinvolved several adjacent sarcomeres.While these particular imaging techniques have not been

used previously to record [Ca2+] transients in cardiac cells,these fluorescent indicators have been utilized photometricallyto record global cellular [Ca2+] transients under these condi-tions (21, 13). Similar to what we observed using imagingmethods, photometry reveals that there is a delay of approx-

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FIG. 2. Determination of intrasarcomeric [Ca2+] gradients in response to electrical stimulation in single cardiac myocytes using fluorescent Ca2+indicators and 3-D digital imaging microscopy. (A) Images of [Ca2+] in a single cardiac myocyte at the indicated times after depolarization usingthe double-pulse protocol. [Ca2+] was calculated from fluorescence of calcium green-2. (C) Images of [Ca2+] calculated from fluorescence of Fura-2at 351 and 380 nm excitation in another single cardiac myocyte. (B and D) Average [Ca2+] along a line drawn parallel to the long axis of the cellin each image inA and C, respectively (4 or 5 ms, long-dashed line, 9 or 10 ms, short dashed line, 14 or 15 ms, solid line). [Ca2+] on the line andin two adjacent voxels (sampled points in a 3-D image) on either side of the line were averaged. (E and F) Surface contour representation of [Ca2+]in the regions delimited by brackets in B and D, respectively.

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imately 5 ms following depolarization before [Ca2+] rises; adelay that presumably reflects the slow activation of dihydro-pyridine-sensitive Ca2+ channels. Between 8 and 12 ms, [Ca2+]rises most rapidly. Hence, the time course of global [Ca2+1transients that emerges from these new imaging methods isquite similar to that obtained by more traditional photometry.

Images obtained by these methods with both dyes reveal thatthe rise in [Ca2+] in response to depolarization is nonuniformand, in places, periodic, with spacing similar to that of thecardiac sarcomere. What is the relation of the nonuniformitiesin the rise in [Ca2+] to the distribution of Ca2+ releaseelements within the sarcomere? In order to assess the rela-tionship of patterns of [Ca2+] gradients to the distribution ofsites of Ca2+ release adjacent to t-tubules, we incubatedmyocytes with Texas Red-labeled wheat germ agglutinin (TR-WGA) to fluorescently label all membranes in contact with themedium. This procedure was previously reported to label boththe sarcolemma and t-tubules (22). Fig. 3 (A and B) showsimages of an intact cardiac myocyte in the presence of TR-WGA. Labeling of t-tubular membranes is clearly evident,although the TR-WGA did not appear to penetrate the fulllength of the t-tubules in the time (10-60 min) typicallyallowed. Fig. 3C shows [Ca2+] transients in relation to theposition of t-tubules in a myocyte in which [Ca2+] was mea-sured with calcium green-2 and the t-tubules labeled withTR-WGA. As can be seen, [Ca2+] increased first and locally

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attained the highest levels throughout the first 15 ms afterdepolarization in the region of the sarcomere containing at-tubule and adjacent junctional sarcoplasmic reticulum (com-pare Fig. SA). A similar correlation between localized [Ca2+]increases and the distribution of the t-tubules was observedwhen [Ca2+] was measured with Fura-2 (Fig. 3D). The corre-lation between regions of the sarcomere exhibiting largest andearliest [Ca2+] increases and the position of the t-tubules wasseen in six out of seven images of six cells in which [Ca2+]increased in a nonuniform and periodic pattern and where theTR-WGA staining pattern was clearly periodic.The observed relation of localized increases in [Ca2+] to the

structure of the sarcomere might well reflect the distributionand time course of Ca2+ release within the sarcomere. How-ever, it is also possible that the pattern of changes in fluores-cence within the sarcomere arises because of differences inoptical properties within the sarcomere. At the near-ultraviolet wavelengths used to excite Fura-2, as much as 20%of fluorescence in Fura-2-loaded cells may be due to cellularautofluorescence. This autofluorescence, while having con-stant intensity for at least 20 ms following stimulation, isprincipally concentrated near the center of the sarcomere. Thisdistribution of autofluorescence could conceivably introduce aspatial bias to the fluorescence signal in Fura-2-loaded cells inresponse to a rise in [Ca2+]. However, it is unlikely that asimilar spatial bias would be introduced into the calcium

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FIG. 3. Relationship of localized [Ca2+] gradients to sarcomere structure. (A) Fluorescence of TR-WGA staining of a single intact cardiacmyocyte. Image shows the projection of the uppermost 18 focal planes of a 3-D image resulting from processing of a series of 38 images taken at0.25-,um intervals through focus. Bottom planes are not shown to facilitate visualization of rows of t-tubules that repeat every 1.8 ,um. (B) Stereopair of images of the bracketed region of the cell inA provide a view from the center of the cell toward the top surface. (C) Relation of [Ca2+Igradients following stimulation of the myocyte to sarcomeric structure. [Ca2+] was calculated from images of calcium green-2 fluorescence. Average[Ca2+] 5, 10, and 15 ms after depolarization along a line parallel to the long axis of the cell is shown by long-dashed, short-dashed, and solid lines,respectively. Variations in TR-WGA fluorescence along that same line (asterisks) indicate the position of t-tubules. (D) Relationship of [Ca2+]gradients following stimulation of another cardiac myocyte to sarcomeric structure. [Ca2+] at 5 (long-dashed), 10 (short-dashed), and 15 ms (solidline) following depolarization in this cell was measured using Fura-2. The position of the z-lines/t-tubules (shown by asterisks) was inferred fromthe distribution of fluorescence measured at 351 nm, a relatively Ca2+-insensitive wavelength. In separate experiments on unstimulatedFura-2-loaded cells, we found that the distribution of 351-nm fluorescence varied within each sarcomere and had its lowest values at positions mostintensely stained by fluorescent TR-WGA. (351-nm fluorescence is thus displayed inverted.) This variation in fluorescence within the sarcomereappears to result from a high density of mitochondria in the middle of most sarcomeres as well as a diminished accessible volume for diffusibledyes of approximately 1000 m.w. near the z-lines.

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green-2 signal, because at the wavelength used to excitecalcium green-2, autofluorescence is barely detectable (2% ofthe average fluorescence signal due to calcium green-2 at 14ms following depolarization). The pattern of [Ca2+] gradientsobserved with Fura-2 and calcium green-2 are quite similar.Therefore it is unlikely that the pattern of [Ca2+] transientsmeasured with Fura-2 is due to a spatial bias introduced by theuneven distribution of autofluorescence within the sarcomere.The pattern of fluorescence changes following stimulation

appears to genuinely reflect spatial patterns in [Ca2+] thatpresumably result from the rapid and local release of Ca2+from the junctional SR and its relatively slower diffusion andsequestration within the sarcomere. If this is indeed the originof the [Ca2+] gradients observed following stimulation, thentreatments expected to alter the rate of release of Ca2+ fromthe SR would be expected to alter the magnitude of the [Ca2+]gradients observed within the sarcomere. By altering the pulseprotocol from a paired pulse stimulation regimen to a singlepulse protocol, the overall [Ca2+] is expected to be reduced byca. 40% (13). Fig. 4A shows [Ca2+] patterns in a single cardiacmyocyte stimulated with both single- and double-pulse proto-cols. Note that while [Ca2+] rose principally in the sameregions of the cell with both protocols, following the single-pulse stimulation, the average [Ca2+] increase was reduced by40 + 8%, and the magnitude of the [Ca2+] gradient as well asthe peak in the [Ca2+] pattern were reduced by 39 + 13%.Coupling between membrane depolarization and Ca2+ releaseis further reduced by run down of Ca2+ currents duringprolonged (>25 min) dialysis of these cells through the patchpipette. When inward Ca2+ currents were lost, neither a [Ca2+]rise nor [Ca2+] gradients were observed following depolariza-tion (data not shown). By contrast, procedures that are knownto increase Ca2+ release from the SR resulted in a larger[Ca2+] rise with larger intrasarcomeric [Ca]2+ gradients. Fig.4B shows the effect of lowering the temperature of a cell from37 to 26°C, a procedure that enhances contraction (7) andincreases global [Ca2+] by approximately 30% (X. Chen andG.I., unpublished results), possibly reflecting the known effectof decreased temperature to prolong ryanodine receptor open-ing (23). As can be seen, while the periodic nature of the [Ca2+]increases was not affected by the drop in temperature, themagnitude of both the [Ca]2+ increases as well as the magni-tude of intrasarcomeric [Ca2+] gradients was increased in thesame cell at the lower temperature. The mean [Ca2+] rise at 15ms was enhanced by 20 + 9%, and the magnitude of the [Ca2+]gradients increased by 16 + 7%. The effect of treatmentsexpected to modulate Ca2+ release on the intrasarcomeric[Ca2+] gradients thus support the notion that they reflectspatial distribution of Ca2+ release sites within the sarcomere.

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FIG. 4. Modulation of [Ca2+] gradients following depolarization byregimes expected to either increase or decrease Ca2+ release from thesarcoplasmic reticulum. (A) [Ca2+] transients in a single myocyte 15ms after depolarization following stimulation by a 1-Hz train of paired(solid) or single (dashed) pulses. [Ca2+] was measured using Fura-2.(B) [Ca2+] transients in a second myocyte in response to depolariza-tion determined at 37°C (solid) and 260C (dashed). Double-pulseprotocol was utilized, and [Ca2+] was measured using Fura-2.

In order to further test this concept, single cardiac myocytes,which were subjected to the same protocol used to assess[Ca>2] transients, were quickly frozen just prior to or 8-15 msafter depolarization, and spatial variations in total [Ca] wereassessed by electron probe microanalysis (14). As summarizedin Fig. SB, during the period when large gradients of [Ca>2]develop in the cytoplasm, the total [Ca] in the junctional SRplummets. This is accompanied by a rise in the total [Ca] in theA-band in regions of overlap of thick and thin filaments. Thetotal [Ca] of the central m-line region does not increasesignificantly. Thus, these results reveal that development of agradient of free cytoplasmic [Ca>2] is associated with releaseof Ca2> from the junctional SR, and its association with regionsof overlapping filaments is presumably due to binding to bothtroponin and myosin.The existence of significant intrasarcomeric [Ca2+] gradi-

ents for at least 15 ms following depolarization differs mark-edly from recent results in skeletal muscle where intrasarco-meric [Ca]2+ gradients dissipated within 4-6 ms (5). Thedifference in the persistence of [Ca2+] gradients is likely to bedue to differences in the time course of Ca2+ release from theSR in these two muscle types. In skeletal muscle fibers, theaction potential is brief (4 ms), and Ca2+ release terminateswith repolarization. In the present studies on cardiac cells,depolarization lasted much longer. Ca2+ must have continuedto be released for at least 15 ms in the cardiac myocytes inorder to account for the continuing rise in [Ca2+] and theincrease in intrasarcomeric [Ca2+] gradients over this interval.

In order to obtain insight into the influence of Ca2+ bindingon the formation of intrasarcomeric [Ca>2] gradients duringCa2+ release, simulations were performed incorporating theeffects of both mobile (Ca2+ indicator) and immobile (myosin,troponin) sites within the sarcomere. This can be seen in Fig.6A and B. The fixed Ca2+ binding sites on the thin and thickfilaments appear to be largely responsible for the steep in-trasarcomere [Ca]2+ gradients by acting as Ca2+ sinks remotefrom sites of release. By contrast, mobile indicators such asFura-2 act to dissipate the [Ca>2] gradient by facilitatingdiffusion of Ca2+ away from sites of release. As can be seen inFig. 6 C and D (boldface lines), the steepness of the [Ca2+]gradient is underestimated by the Fura-2 fluorescence ratiodue to limits imposed by the kinetics of interaction of Ca2+with Fura-2. The gradient in fluorescence ratio is furtherunderestimated because of blurring even with the partial

A

B Regional changes in total [Ca] in single cardiac mXrocvtesI-band (junct. SR) A-banid (myofilamcnts) m-line

Diastoic 2.30-h.10 (n=12)So stole (0.15--0.04 ini22)

0.61±-0.04 (n=30) 0.,-L0. 06 (n=l )(1.86-0.03 (n=34) 0.35-0.03 (n=20)

FIG. 5. Electron probe microanalysis of the spatial variation oftotal calcium concentration, [Ca], in cardiac myocytes quickly frozenwhile under voltage clamp. (A) Transmission electron micrograph ofa guinea pig trabecular myocyte quick frozen in a manner similar toprocedures used in B for analysis of [Ca] distribution. Specimen wasfreeze-substituted, embedded, and stained conventionally (21). t,t-tubule; jSR, junctional SR; m, mitochondrion; Z, z-line; H, m-line;scale bar, 0.5 ,um. (B) Total [Ca] distribution in mM, in sarcomeres ofsingle myocytes quickly frozen either in diastole or 8-15 ms after startof depolarization following a train of pulses. n equals the number ofsarcomeres analyzed. Myocytes frozen for electron probe microanal-ysis were neither freeze-substituted nor stained or embedded.

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Proc. Natl. Acad. Sci. USA 93 (1996)

0 0.2 0.4 0.6 0.8distance from z-line (um)

0 0.2 0.4 0.6 0.8distance from z-line (um)

FIG. 6. Simulation of the spread of Ca2+ and its detection byoptical methods within the sarcomere of a cardiac myocyte followingdepolarization. (A) Distribution of [Ca2+] 3 ms after onset of Ca2+release at the z-line assuming neither Ca2+ binding to troponin andmyosin or Fura-2 (solid line), with Ca2+ binding to troponin and myosin(dashed line), and with binding both to myosin, troponin, and Fura-2(boldface line). (B) Same as inA, but 15 ms after onset of Ca2+ release.(C) Distribution of ratio of fluorescence at 351/380 within a sarcomere3 ms after the onset of Ca2+ release with binding to troponin, myosin, andFura-2 but without blurring introduced by microscope optics (boldfaceline), with blurring introduced by microscope optics (dashed with circles),and with blurring introduced by microscope optics and partial reversal ofthat blurring by processing with constrained deconvolution algorithm(dot-dashed). (D) Same as C but 15 ms after onset of Ca2+ release.

confocal illumination, which reduces out of focus light; theblurred gradient in fluorescence ratio is 71% of the value of themodel (at 3 ms). Deconvolution improves our estimate of thisgradient of fluorescence ratio to 88% of the value of the model(at 3 ms). Finally it is likely that the [Ca2+] reached in thelimited 12 nm space between the t-tubule and Ca2+ releasesites of the junctional SR is considerably higher than what wehave reported or even modeled. The size of the distance stepsused in the model (30 nm) and the resolution achieved afterdeconvolution of our images are likely too coarse to accuratelycalculate the [Ca2+] in this physiologically important but verysmall "fuzzy space" (24).The direct demonstration of large intrasarcomeric [Ca2+]

gradients during the onset of the excitation contraction cou-pling in cardiac muscle provides support for earlier theoretical(25) and experimental (26, 27) work, which suggested thatCa2+ release is initiated at the junctional SR. The existence oflocally high [Ca2+] near the t-tubules and junctional SR duringthe onset of activation may explain why a number of Ca2+-sensitive processes in that region are activated, although [Ca2+]around the myofibrils has not yet reached levels expected toinfluence these Ca2+-sensitive processes (24). For example,termination of SR Ca2+ release following depolarization maybe brought about, at least in part, by the observed large localincrease in [Ca2+] in the t-tubular/junctional SR region. Thiswould be expected to decrease Ca2+ release both by decreasingthe driving force on Ca2+ and by interacting with the ryanodinereceptor to inactivate the channel (28). The development ofthis locally high [Ca2+] in the cytoplasm adjacent to thet-tubule in the first 15 ms after depolarization may also explain,at least in part, why as early as 20 ms after the onset of thecardiac action potential Ca2+ movement across the t-tubules

reverses direction (29). The local high [Ca2+] that developswithin this interval would be expected to cause a reversal in thedriving force of the Na+/Ca2+ exchanger, which is concen-trated in the t-tubules (30). The variation in [Ca2+] transientsobserved between groups of sarcomeres may reflect regionaldifferences in both the extent of Ca2+ loading ofjunctional SRas well as the local availability of L-type Ca2+ channels (21).Such differences, which appear to involve areas containingseveral sarcomeres, may in turn reflect local variations in cyclicAMP-dependent protein kinase activity, which is known toaffect these processes (31, 32). While the basis for suchregional differences is not known, the effect will be that forceby sarcomeres experiencing different [Ca2+] will vary. Owingto the ascending nature of the length-tension relationship incardiac muscle (7), however, these regional differences will notbe expected to induce mechanical instability for the cell.

We thank Doug Bowman for help in development of specialsoftware for these experiments, Jeff Carmichael for photographicassistance, and Debra Roy for secretarial assistance in the preparationof this manuscript. This work was supported in part by grants GM14157(E.F.E.) and HL14523 (F.S.F.) from the National Institutes of Health,BIR-9200027 (F.S.F.) from the National Science Foundation, and theNorth Atlantic Treaty Organization.

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Nature (London) 367, 739-741.6. Winegrad, S. (1965) J. Gen. Physiol. 48, 997-1002.7. Bers, D. M. (1991) Excitation-Contraction Coupling and Cardiac Con-

tractile Force (Kluwer Academic Publishers, Dordrecht).8. Otsu, K, Willard, H. F., Khanna, V. K, Zorzato, F., Green, N. M. &

MacLennan, D. H. (1990) J. Biol. Chem. 265, 13472-13483.9. Fabiato, A. (1985) J. Gen. Physiol. 85, 189-320.

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11. Cheng, H., Lederer, W. J. & Cannell, M. B. (1993) Science 262,740-744.12. Lopez-Lopez, J. R., Shacklock, P. S., Balke, C. W. & Wier, W. G.

(1995) Science 268, 1042-1045.13. Han, S., Schiefer, A. & Isenberg, G. (1994)J. Physiol. 480, 411-421.14. Wendt-Gallitelli, M. F. & Isenberg, G. (1991)J. Physiol. 435,349-372.15. Carrington, W. A., Lynch, R. M., Moore, E. D. W., Isenberg, G.,

Fogarty, K. E. & Fay, F. S. (1995) Science 268, 1483-1487.16. Grynkiewicz, G., Poenie, M. & Tsien, R. Y. (1985) J. Biol. Chem. 260,

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A. J. (1991) J. Physiol. 434, 469-488.24. Lederer, W. J., Niggli, E. & Hadley, R. W. (1990) Science 248, 283.25. Wohlfahrt, B. & Noble, M. I. M. (1992) Pharnacol. Ther. 16, 1-43.26. Wendt-Gallitelli, M. F., Jacob, R. & Wohlburg, H. (1982) Z. Natur-

forsch 37, 712-720.27. Jorgensen, A. O., Shen, A. C.-Y., Arnold, W., McPherson, P. S. &

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(1987) Circ. Res. 61, 17-23

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