Pacing rate, halothane, and BDM affect fura 2 reporting of [Ca2+]i in intact rat trabeculae

Yandong Jiang and Fred J. Julian

Department of Anesthesia Research Laboratories, Brigham and Women's Hospital, Boston, Massachusetts 02115

    ABSTRACT
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Abstract
Introduction
Methods
Results
Discussion
References

Experiments were done on intact trabeculae from rats. Fura 2 in the salt form was microinjected directly into the myoplasm. The experiments were conducted at 30°C, with 2 mM extracellular Ca2+ concentration and pacing at either 0.5 or 5 Hz. The aims were to establish a new method for in vivo calibration of fura 2 and to determine the effect of autofluorescence changes on intracellular Ca2+ concentration ([Ca2+]i) reported by fura 2. Autofluorescence was recorded under optimal conditions for fura 2 fluorescence (emission at 510 nm). By alteration of the oxidation-reduction state, it was shown that NADH is the main component of autofluorescence in heart. An increase in pacing frequency caused a decrease in autofluorescence. Both halothane and 2,3-butanedione monoxime (BDM) at 5-Hz pacing produced a substantial rise in autofluorescence, approaching the levels observed at 0.5-Hz pacing. The values for the dissociation constant (678 nM) and maximum fluorescence ratio of fura 2 for Ca2+ for the in vivo calibration are 3.4 times larger and 2.6 times smaller, respectively, than those found in vitro. Using the parameters obtained in vivo, we found that the diastolic and systolic [Ca2+]i of a twitch at 30°C were 0.2 and 2.4 µM, respectively. Proper correction of the autofluorescence change unmasks the [Ca2+]i elevation caused by 5-Hz pacing. It was concluded that autofluorescence is not constant and that interventions affecting autofluorescence need correction if fura 2 is used to report [Ca2+]i.

heart; cardiac; fluorescence; intracellular calcium concentration; volatile anesthetic; 2,3-butanedione monoxime

    INTRODUCTION
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Abstract
Introduction
Methods
Results
Discussion
References

FURA 2 HAS BEEN USED WIDELY to report intracellular Ca2+ concentration ([Ca2+]i) since it was first introduced (17). A novel technique was developed by which the polyanionic salt form of fura 2 was directly loaded into the myoplasm of a single myocyte, from which it spread throughout an entire trabecula via the gap junctions (3). This technique is an advance over the acetoxymethyl ester of fura 2 (fura 2-AM) loading, which causes accumulation of fura 2 into other intracellular compartments or organelles such as the sarcoplasmic reticulum (SR) or mitochondria (30). It also avoids the complications resulting from incomplete hydrolysis of fura 2-AM (28). With the use of fura 2, [Ca2+]i is determined by the ratio (R) of the fluorescence of the dye for two excitation wavelengths. This makes it possible to measure [Ca2+]i independently of dye concentration, path length, excitation intensity, absolute instrumental sensitivity, and photobleaching (32). In addition, use of the R signal reduces motion artifact during contraction and relaxation. The fluorescence used to form R must be dye-related fluorescence only, and any other fluorescence present must first be subtracted. Background fluorescence is easy to correct for because it remains essentially constant; however, tissue autofluorescence can be large and variable (1, 19, 31). If the autofluorescence change is not properly accounted for, it could result in an inaccurate estimation of R, and therefore [Ca2+]i, using fura 2.

In some mammalian tissues, like the heart, autofluorescence is intense (1, 19) and is caused largely by the reduced pyridine nucleotide (NADH) in mitochondria (12). The strong NADH-dependent autofluorescence in cardiac muscle can hardly be considered constant because it varies, for example, with pacing protocol (8) and oxygenation (10). It is also sensitive to treatment with various agents such as the well-known vapor anesthetic agent halothane, which causes autofluorescence (from NADH) in the heart to increase (22). An incorrect conclusion could be reached that a directional change in R caused by an autofluorescence change signaled a similar directional change in [Ca2+]i, whereas the opposite might be true because of concurrent changes in autofluorescence.

Accuracy of intracellular Ca2+ measurement depends on a reliable calibration method for relating R to [Ca2+]i. Even though directly loading fura 2 salt into the myoplasm can overcome many drawbacks of fura 2-AM loading, as mentioned above, there are still several factors complicating in vivo calibration. First, the properties of fura 2 are altered from those in solution, as a result of the actions of various factors such as ionic strength, viscosity, and protein concentration (32). Second, the [Ca2+]i may not reach equilibrium with the extracellular Ca2+ concentration ([Ca2+]o) during the calibration because of membrane potential-dependent and transporter processes acting in the surface membrane to oppose equilibration. Finally, autofluorescence may be dramatically altered during calibration. Therefore, it is now generally accepted that calibration of fura 2 in aqueous media is unreliable for use in determining [Ca2+]i. This has led to the use of various in vivo protocols developed by different workers, although generally Ca2+ ionophores are used to clamp [Ca2+]i to various levels followed by measurement of R (21). Recently, another calibration method for myocytes, which takes advantage of the powerful Na+/Ca2+ exchanger in heart to equilibrate extra- and intracellular [Ca2+], has been presented (4). With use of this method, incomplete equilibration of extra- and intracellular [Ca2+] is no longer a problem. However, little attention has been directed to autofluorescence changes during an in vivo calibration. A commonly used procedure of in vivo calibration in heart muscle has been developed, called the metabolic inhibition method (23). In this method, myocytes are first ATP depleted in a glucose-free medium containing an inhibitor and uncoupler of oxidative phosphorylation and then treated with buffered solutions of Ca2+ with ionophores to allow estimation of R as a function of [Ca2+]i, assuming unchanged autofluorescence. However, this method does not seem to take account of changes in NADH content, and it has been shown that various agents that influence oxidative phosphorylation in the mitochondria can greatly affect the level of autofluorescence (8). Therefore, these effects, if not taken into account, could produce incorrect calibration curves, thus leading to inaccurate determination of [Ca2+]i.

It was one aim of these experiments to clarify the experimental conditions under which autofluorescence in the heart can be considered relatively constant, making it possible to assume that the level of autofluorescence before fura 2 loading can be used as an autofluorescence correction throughout the experiment. Another aim was to find a reliable calibration technique for use to relate R to [Ca2+]i in isolated multicellular trabeculae and thus make it possible to determine accurately diastolic and systolic [Ca2+]i. We found here that autofluorescence changes, if not properly corrected for, could significantly affect the determination of [Ca2+]i during both an experiment and a calibration.

    METHODS
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Methods
Results
Discussion
References

Dissection procedure. Male Lewis-Brown-Norway rats (150-175 g) were used in these experiments (Harlan, Indianapolis, IN). The procedure described was approved by the Harvard Animal Care and Use Committee. Rats were given intraperitoneal injections of calcium heparin (400 U), and 20 min later deep anesthesia was produced by intraperitoneal injections of pentobarbital sodium (60 mg/kg). Hearts were quickly removed and placed in a cold modified Krebs solution (MKS). The hearts were then perfused retrograde through the aorta using a Langendorff apparatus for 10 min at 22°C with the same MKS, containing (in mM) 108 NaCl, 6 KCl, 1.2 MgCl2, 2 CaCl2, 24 NaHCO3, 10 sodium pyruvate, and 4 glucose and 5 U/1,000 ml insulin and were bubbled with 95% O2-5% CO2, final pH 7.4. Finally, the hearts were flushed with dissection solution made by addition of 15 mM KCl to the standard MKS (final KCl concentration = 21 mM) while they were still mounted on the Langendorff apparatus.

A heart was pinned in a dish filled with dissection solution, and the right ventricular wall was dissected under microscopic observation along both sides of its juncture with the interventricular septum, thereby exposing the inner surface of the right ventricle. Trabeculae similar to those described previously (14, 26) were isolated. These muscles (length ~2.5-4 mm) can be identified as originating from a cusp of the right atrioventricular (tricuspid) valve and terminating on the interventricular septum or right ventricular wall. Suitable long, thin, nonbranching trabeculae were dissected for experiments, and these were thin enough to allow microscopic observation of striation patterns. The very small size of the preparations (thickness of ~50 µm) also obviates the concern about adequate oxygenation in a cardiac muscle isolated from normal blood perfusion (29). Monofilament nylon surgical suture (Ethilon 10-0; Ethicon, Somerville, NJ) was used to tie a small piece of valve attached to one end of each trabecula to a force transducer (model 400A; Cambridge Technology, Cambridge, MA), and the other end was tied to a stainless steel wire (0.2 mm diameter) connected to a three-dimensional micropositioner. Cyanoacrylate tissue adhesive (Histoacryl blau; B. Braun, Melsungen, Germany) was used to fix firmly the two ends of the preparation to the force transducer wire and micropositioner wire. This method of attachment considerably decreases extra end compliance (20).

Bath setup and electrical stimulation. The trabecula was put into a temperature-regulated chamber (8 × 6 × 5 mm) milled into a Lucite plate 6 mm thick. The chamber had a glass coverslip floor (thickness 0.12 mm) to facilitate near-ultraviolet light transmission. The Lucite plate with its chamber was positioned on the stage of an inverted microscope (Nikon Diaphot 300) fitted with a fluorescence system [Photon Technology International (PTI) Deltascan 4000]. Trabeculae were superfused at a rate of 12 ml/min with MKS bubbled with 95% O2-5% CO2. Two semi-closed bottles (standard and treatment) were placed on a shelf ~2 ft above the level of the chamber, thus allowing the use of a gravity-driven flow system. The MKS solutions in the bottles were vigorously bubbled with 95% O2-5% CO2. The bottle outputs were connected through a simple Y arrangement, with a stopcock switch allowing preparations to be superfused with contents from either bottle. When required, 2,3-butanedione monoxime (BDM; Aldrich, Milwaukee, WI) solutions were obtained by adding the desired amounts directly to MKS in the treatment bottle, as was the halothane (see Halothane below). The temperature in the bath was maintained at 30 ± 0.2°C using two in-line heaters in series (model TC-324A; Warner Instrument, Grand Haven, MI). The preparations were stimulated at 0.5 or 5 Hz with 5-ms pulses via platinum plate electrodes in the bath that were connected to a stimulator (model S88; Grass Instruments, Quincy, MA) driven by an interval generator (model 830; World Precision Instruments, New Haven, CT) and pulse module (model 831A, World Precision Instruments). The stimulus strength was adjusted to 1.5 times threshold, and tension was continuously monitored on both a chart recorder (model 7133A; Hewlett-Packard, Palo Alto, CA) and one channel of a digital oscilloscope (Nicolet 4094B). The optimal tension response with respect to length was obtained by stretching the trabecula using the three-dimensional micropositioner attached to one end. A TV camera (model HV7200, Hitachi) receiving long-wavelength light (>620 nm) from the side port of the microscope made it possible to observe the striation pattern under red light transillumination on a video monitor (model WV5410, Panasonic). Resting sarcomere length resulting in maximum active force generation was in the range of 2.18 to 2.30 µm. These values were measured from hard copy striation patterns sampled from the video image (using a Nikon ×40 objective) with a UP1200 Mavigraph color video printer (Sony).

Fluorescence system. The fluorescence was collected using either the photon-counting or the analog mode of the PTI Deltascan 4000 system. A 75-W xenon lamp was the source of excitation light. The light passed through monochromators (entrance and exit slit widths set to 2 nm for photon-counting mode and 10 nm for analog mode) that were programmed to select wavelengths of 345 and 380 nm for excitation of fura 2. When twitches were recorded, the wavelength used was toggled at the stimulating frequency so that, during successive twitches, the muscle was illuminated by alternate wavelengths. Light exiting the monochromator was carried by a quartz fiber-optic bundle to a filter cassette coupled to the microscope objective, which also functioned as a condenser lens. Upon entering the cassette, the light was first short-pass filtered (<470 nm) to remove most of the residual light that could interfere with the collection of fura 2 fluorescence before reflecting off a dichroic mirror (400 nm long pass) en route to the objective. In these experiments, we used an Olympus (Lake Success, NY) ×10 (Dapo UV/340; numerical aperture, 0.4) objective, which gave good transmission at wavelengths >340 nm. Fluorescence from the preparation passed back through the objective and dichroic mirror to the side port of the microscope. Here, a second dichroic mirror passed long-wavelength light (>620 nm) to the TV camera and shorter wavelengths to a photometer with a rectangular region of interest (ROI) adjustment that could be viewed through an eyepiece when measurements were not being made. The ROI defined the boundaries in the object plane, from which light emitted from the preparation was collected and sent to two photomultipliers. The current from one photomultiplier, receiving 92% of the light, was amplified and filtered to produce an analog signal, which was used for dynamic recording. The current from the other photomultiplier was used for photon counting, and this mode was used to record wavelength scan spectra and monitor autofluorescence. Just before reaching each of the photomultiplier tubes, light signals were passed through emission filters that transmitted wavelengths of 510 ± 20 nm, these being near the peaks of the emission signals for Ca2+-free and Ca2+-bound fura 2. The voltage output from the analog photomultiplier was sent to a low-pass analog filter with a cutoff frequency of 500 Hz (model SR650; Stanford Research Systems, Sunnyvale, CA) and then to a four-channel oscilloscope (Nicolet 4094B) sampling at a frequency of 2 kHz. A dual-disk recorder (Nicolet XF-44/2) was used to record to floppy disks the tension and fluorescence signals acquired by the digital oscilloscope. For wavelength scan spectra, the fluorescence signal at 510 nm was measured as the excitation wavelength was scanned over a range of values (300-450 nm). This made it possible to measure dye loading as well as the influence of other treatments on fluorescence properties. The system was controlled by PTI software ("Oscar") running on an IBM-compatible 486 computer (American Megatrends) that could be used to control shutter openings, monochromator settings, and other automated data acquisition programs.

Dye loading procedures. Before the trabeculae were loaded with fura 2, autofluorescence (emission at 510 nm) was measured, first by scanning the excitation wavelengths from 300 to 450 nm and then by taking individual analog measurements at excitations 345 and 380 nm at a pacing rate of 0.5 Hz and a temperature of 30°C. Preparations were then loaded by iontophoretic injection of fura 2 dye, by a method already described (3), in MKS with [Ca2+]o of 0.5 mM at 22°C with no stimulation. Micropipettes were pulled on a Flaming/Brown micropipette puller (model P87; Sutter Instrument, Novato, CA), and tips were filled with 1 µl of 2 mM fura 2 (pentapotassium salt, Molecular Probes, Eugene, OR). The pipettes were then back filled with a 150 mM potassium acetate solution (pH = 7.3) and connected via a pipette holder to the headstage of a World Precision Instruments electrometer module (model S-7071A), which permitted the measurement of membrane potential or electrode impedance and had the option to pass currents of varying amplitude. Tip resistance of 200-300 MOmega was typical in micropipettes filled with the fura 2 solution before impalement of single myocytes, and stable membrane potentials of -40 to -60 mV were required for successful dye loading. After impalement, injection of the dye into the myocyte was achieved by passing a negative current of 3-5 nA, which resulted in a 15- to 25-mV hyperpolarization of the cell. Dye loading could be monitored through the binocular eyepiece of the microscope. A bright region of fluorescence could be seen emanating from the point of impalement when light of an excitatory wavelength (380 nm) was focused on the preparation. After loading, the preparation was superfused again with the standard MKS, the temperature was increased to 30°C from room temperature (~22°C), and the pacing was resumed at 0.5 Hz. In agreement with earlier results (3), the initial localized bright region of fluorescence gradually spread throughout the illuminated part of the muscle so that by the time of data recording ~1 h later, complete uniformity of fluorescence was present. Fura 2 was loaded to levels such that the added fluorescence was at least three times the autofluorescence observed at 358 nm (the isosbestic point for fura 2) before loading.

Halothane. When required, halothane was added to the superfusion solution in the treatment bottle. A Dräger vaporizer was set to deliver a concentration (vol/vol) in the treatment bottle sufficient to produce a repeatable and reversible attenuation of twitch-developed force by ~90%. The resulting concentration of halothane in the MKS was near 0.5 mM, as determined using a Hewlett-Packard 5890 series II gas chromatograph with a Hewlett-Packard 624 column and a flame ionization detector. According to a recent report, the 1 minimal alveolar concentration (MAC) of halothane in non-blood-containing aqueous medium at below normal body temperature in the rat should be ~0.27 mM (15), so our dose is nearer to 2 MAC. The MAC concept is related to an intact whole body response to a noxious stimulus, signaled by movement in 50% of the subjects, so that MAC values near 2 are still clinically relevant to the study of mechanisms of action of anesthetic agents.

Calibration. The solutions for the calibration of fura 2 are similar to those described elsewhere (4). There are two solutions, A and B. Solution A contained (in mM) 15 NaCl, 135 KCl, 1.5 MgCl2, 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid, 10 glucose, 10 caffeine, 10 BDM, 10 ethylene glycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA), 0.03 2,5-di(tert-butyl)-1,4-benzohydroquinone (TBQ), 0.05 strophanthidin, and 0.05 A-23187 (Ca2+ ionophore). Solution B contained all of the components of solution A together with 10 mM CaCl2. The pH of both solutions was adjusted to 7.2 with KOH at 30°C.

The in vitro calibration was performed in the experimental chamber without trabeculae at 30°C. Fura 2 (5 µM) was added to the calibration solutions. The minimum fluorescence ratio (Rmin) was obtained using solution A, whereas the maximum fluorescence ratio (Rmax) was measured in solution B, which contained, in addition, 1 mM CaCl2, so that the total [Ca2+] was 11 mM. Intermediate [Ca2+] were obtained by mixing appropriate volumes of solutions A and B. Each solution was replaced by a solution with a higher [Ca2+] until the solution for Rmax was reached. The [Ca2+] in the various solutions was determined using a Ca2+ electrode (Orion, model 93-20) and a previously described technique (6). The values for the ratio of Ca2+-free to Ca2+-bound fluorescence at 380 nm (beta ) were obtained from the fluorescence excited at 380 nm by dividing the fluorescence signal in the Rmin solution by that in the Rmax solution. Values for the dissociation constant (Kd) were obtained as described in RESULTS.

For the in vivo calibration, which was also done in the experimental chamber at 30°C, the trabeculae were loaded with fura 2 as described above at 22°C until the dye fluorescence (at isosbestic 358 nm) was ~10 times that of the combined autofluorescence and background fluorescence. This degree of fura 2 loading subsequently caused the twitch and Ca2+ transient signal to be markedly attenuated. After the dye was evenly distributed through the whole preparation, 10 mM BDM was added to the MKS to minimize injury that could be caused by a subsequent caffeine-induced contracture when the MKS was replaced by solution A. After this, the R signal gradually decreased and leveled off after ~10 min as it reached Rmin. After Rmin had been recorded, solution A was replaced sequentially by the various mixtures of solutions A and B to produce a range of free Ca2+ concentrations. Each solution was replaced by a solution with a higher [Ca2+] until the solution for Rmax was reached. At each [Ca2+], an equilibration time of 20 min was allowed, thus taking ~100 min to complete the series of measurements from Rmin to Rmax. The [Ca2+] were determined as for the in vitro case. The fluorescence intensity excited at 358 nm was obtained by wavelength scan in the initial Rmin solution and in the final Rmax solution. It was found that the isosbestic fluorescence decayed at an average rate of ~23%/h at 30°C, but this high decay rate was mainly because of Rmax solution. As above, the value for beta  was obtained using the fluorescence intensity excited at 380 nm in the Rmin solution divided by that in the Rmax solution. Before the division for beta , the fluorescence intensity (F) excited at 380 nm in the Rmax solution was corrected for dye loss by multiplying by the isosbestic ratio F358, Rmin/F358, Rmax.

Data analysis. Fluorescence and tension records were analyzed using Vu-Point3 data analysis software (Maxwell Laboratories, San Diego, CA) and SigmaPlot 4.0.

    RESULTS
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Methods
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It has been mentioned above that fluorescence unrelated to fura 2 consists of two parts: background fluorescence and tissue autofluorescence. These must be summed and subtracted from the total fluorescence after loading fura 2 before forming the ratio R. It is straightforward to measure the essentially constant background fluorescence; however, tissue autofluorescence in the heart can vary depending on experimental conditions. Therefore, under optimal conditions for fura 2 fluorescence measurement, it is important to know the excitation wavelength distribution of the autofluorescence as well as to determine what causes it. Figure 1 shows a typical excitation wavelength scan in a rat trabecula before and after loading fura 2. Typically, the fluorescence intensity after loading was ~3 times that present initially, which is close to that previously reported (3). It is apparent in Fig. 1 that the fura 2-loaded scan overlaps very closely the wavelength domain of the autofluorescence scan before loading. If the autofluorescence measured under the conditions optimal for fura 2 and not for NADH is mainly because of NADH, it could be variable and interfere with estimation of dye fluorescence and, consequently, R. Therefore, experiments were done to test whether increases or decreases in the intracellular concentration of NADH ([NADH]i) affect the autofluorescence overlapping with that of fura 2, and the results are shown in Fig. 2. The autofluorescence in the absence of fura 2 was markedly reduced by carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP), which causes a decrease in [NADH]i by uncoupling oxidation-phosphorylation, and was greatly increased by cyanide, which causes an elevation in [NADH]i by inhibition of the respiratory chain (8). FCCP and cyanide, respectively, caused a decrease of 85% and an increase of 105% in autofluorescence at 358-nm excitation compared with the control value. The resulting excitation scan wavelength spectra were subtracted from the controls to form difference spectra. These difference spectra were then compared with the spectrum of a solution of NADH (1). The results are shown in Fig. 2, A and B for FCCP and in C and D for cyanide. Particularly for FCCP, there is a very close correspondence between the difference and NADH spectra, whereas, in the case of cyanide, the agreement is slightly less than perfect. It is possible that other tissue components, such as riboflavin and flavin coenzymes (1), also contribute slightly to autofluorescence. The appropriate rise and fall of autofluorescence in response to substances known to change the level of NADH, together with the spectral similarity, strongly suggest that most of the autofluorescence present in isolated, intact trabeculae comes from NADH, even in a microscope system optimized for fura 2 recording. These results emphasize that there is a strong tissue autofluorescence in heart muscle and that there is overlap between the excitation wavelength scans of the autofluorescence and fura 2. Therefore, control experiments are required to determine the effect of interventions on autofluorescence in the absence of fura 2. 


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Fig. 1.   An excitation wavelength scan spectrum from a trabecula recorded at an emission wavelength of 510 nm before and after loading of fura 2. Isolated trabecula obtained and mounted as described in METHODS, with temperature 30°C and pacing frequency 0.5 Hz. The trace labeled autofluorescence (corrected by subtracting out background fluorescence) shows the typical tissue autofluorescence distribution of an isolated trabecula before loading with fura 2 salt. Note that autofluorescence is near maximal at ~355 nm, with strong contributions at both 345 and 380 nm. After fura 2 loading, fluorescence intensity was increased, with peak remaining near 355 nm. Thin vertical lines have been drawn in plot, with origins on abscissa at 345 and 380 nm. Each trace is an average of 5 wavelength scans, with wavelength step size of 1 nm and integration time of 0.05 s.


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Fig. 2.   Excitation wavelength scan spectra from trabeculae of autofluorescence at emission 510 nm before and after treatments with carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) or cyanide. Effects on autofluorescence of treatment with FCCP are shown in A and B. A: curve labeled control is from an untreated trabecula, whereas curve labeled FCCP is from an FCCP-treated trabecula (10 µM) to show markedly diminished autofluorescence intensity. B: difference spectrum of control minus FCCP from A is normalized to excitation scan of a solution of 100 µM NADH. Note very similar wavelength distribution, suggesting that a decrease of NADH content is responsible for change in autofluorescence. In C and D, effects on autofluorescence of treatment with cyanide are shown. C: curve labeled control is from an untreated trabecula that was then treated with cyanide (5 mM) to produce curve labeled cyanide. Cyanide markedly increased autofluorescence intensity. D: difference spectrum of cyanide minus control from C is normalized to excitation scan of a solution of 100 µM NADH, with both curves scaled as above. Two curves have a similar wavelength distribution, suggesting that, to a large extent, the effect of cyanide treatment shown in C is to increase NADH content of trabecula.

It is likely that there may be substantial changes in NADH level in trabeculae, depending on the pacing protocol, temperature, oxygenation, substrate supply, and presence of any agents capable of affecting the NADH level. An example of the influence of pacing frequency is shown in Fig. 3 in a trabecula not loaded with fura 2. Here, all other parameters except pacing frequency were kept constant and the temperature was 30°C. In A, the raw autofluorescence at 510 nm excited by 345-nm light is shown, whereas in B the twitch force output associated with the 345-nm trace is also shown on the same time base. The pacing frequency was gradually increased from 0.5 to 5 Hz (30 to 300 counts/min). As the pacing frequency increased, the autofluorescence intensity at 345 nm dropped but rose rapidly to a higher level near the initial when the stimuli were turned off. The entire process reversed itself as the pacing frequency was decreased. The twitch force response is noteworthy because it shows an opposite reaction to that of the autofluorescence, thus revealing a positive force-frequency relationship.


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Fig. 3.   Single wavelength measurement of autofluorescence intensity and corresponding twitch force at different pacing frequencies. Autofluorescence was excited at 345 nm, and emission was collected at 510 nm. Photon counting mode was used with a sampling rate of 200 counts/s. Temperature was 30°C. Autofluorescence intensity levels (A) and corresponding twitch forces (B) are shown on same time base. Pacing frequency, indicated in A and B by number above each segment, was incremented from 0.5 Hz to 1, 2, 3.3, and 5 Hz, and this was followed by a return to initial condition along same path from 5, 3.3, 2, 1, and finally 0.5 Hz. Zeros indicate segments with no stimulation. Diastolic autofluorescence intensity from center of each pacing segment (measured using a greatly expanded time base) together with averaged fluorescence intensity for entire segment were obtained from A and B, and these are plotted against appropriate pacing frequency in C. At highest pacing frequency (5 Hz), 3 sample points could be obtained during diastole when force had returned to baseline. Filled symbols are diastolic autofluorescence: circles, increasing Hertz; triangles, decreasing Hertz. Open symbols are averaged autofluorescence: squares, increasing Hertz; diamonds, decreasing Hertz. At certain Hertz, e.g., 2 and 3.3, values are so close that some symbols are hidden.

The single wavelength-determined decrease in autofluorescence as pacing rate is increased could be a spurious result from a motion artifact caused either by translation or by change in average position or orientation of the muscle. Our use of a sampling rate of 200 counts/s allowed us to extract from the data shown in Fig. 3A both the diastolic and averaged autofluorescence intensity from each of the segments associated with a particular pacing rate. The results are shown in Fig. 3C as a function of pacing frequency. It is apparent that both diastolic and averaged autofluorescence decrease very similarly with increasing pacing frequency. This means that the autofluorescence from a resting muscle within a given pacing-frequency segment is not much different from the averaged autofluorescence over the entire pacing-frequency segment. Thus contraction-induced changes per se in autofluorescence cannot account for the major part of the decline in autofluorescence with increasing pacing frequency. This finding argues in favor of the view that pacing frequency, not effects associated with motion artifact, is the most important parameter influencing autofluorescence. Somewhat similar results have been presented (7).

An important question concerns whether the use of various agents in the course of an experiment might influence the NADH level and thus affect the use of fura 2 to report [Ca2+]i. One agent of particular interest to us is the common vapor anesthetic halothane, which has been reported to cause a dose-dependent increase in NADH fluorescence in the heart (22). Experiments were done in the absence of fura 2 loading to test for this possibility, and the results are shown in Fig. 4, A and B. The raw autofluorescence intensities produced by 345- and 380-nm excitation are shown as a function of time while the trabecula was being paced at 0.5 Hz (A) and 5 Hz (B). Halothane at a dose sufficient to attenuate twitch force reversibly by approx 90% (not shown) was added to and then removed from the bathing medium. It is evident that the effect of halothane was an essentially completely reversible increase in autofluorescence, which was substantial only in the case of pacing at 5 Hz. This is in agreement with earlier work reported for isolated rat hearts that were kept at 37°C and also paced at 5 Hz (22). The important parameter to consider, as shown in Fig. 4, is the pacing frequency. When the pacing frequency is high, such as 5 Hz, the autofluorescence is diminished in the absence of halothane, as shown in Fig. 3, so that the halothane effect, essentially a reversal of the high pacing rate effect, is significant. However, when the pacing frequency is low, such as 0.5 Hz, our standard rate, autofluorescence is high and the consequent rise induced by halothane is much smaller. Another result is also shown in Fig. 4, concerning the use of BDM, a potent inhibitor of active force generation and a blocker of attached, strongly bound cross bridges (2, 24). As in the case of halothane, only the pacing frequency was varied: 0.5 Hz (C) and 5 Hz (D), whereas the dose of BDM depressed active force generation by ~95% in both C and D (not shown). There is a striking similarity of results between halothane and BDM. In each case, a strong effect on autofluorescence is only observed at the higher pacing frequency.


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Fig. 4.   Autofluorescence intensity at emission 510 nm and excitation wavelengths 345 and 380 nm. Pacing frequency was either 0.5 or 5 Hz, with and without treatment with either halothane or 2,3-butanedione monoxime (BDM) at 30°C. Fluorescence was collected in photon-counting mode at a collection rate of 200 counts/s. Dose of halothane reduced steady-state twitch force amplitude by ~90%. Halothane treatment period is between arrows at 30 and 270 s. In A, at 0.5 Hz pacing, there was minimal influence on autofluorescence level. However, in B, when pacing frequency was increased to 5 Hz, there was a marked increase in autofluorescence caused by halothane, particularly in the 345-nm trace. Halothane effects on autofluorescence and twitch amplitude were reversible. Influence of BDM (20 mM) on autofluorescence of an isolated trabecula is also shown: 0.5 Hz (C) and 5 Hz (D). Fluorescence was collected as described above for halothane. Dose of BDM reduced steady-state twitch force amplitude by ~95%. BDM was applied during same time period as for halothane. At 0.5-Hz pacing, as with halothane, there was minimal influence on autofluorescence levels. However, when pacing frequency was increased to 5 Hz, there was a marked increase in autofluorescence caused by BDM, particularly in the 345-nm trace. BDM effects on autofluorescence and twitch amplitude were reversible.

Although the conditions under which the dye fluorescence ratio R can now be reliably estimated (low pacing frequency, etc.), an accurate transformation from R to [Ca2+] still remains to be found. To show the need for an in vivo calibration, we conducted both in vivo and in vitro calibrations under nearly identical conditions at 30°C. An in vitro calibration of fura 2 was carried out in the calibration buffer solutions containing 5 µM fura 2 and free Ca2+ ranging from ~1 nM to >1 mM. The results are shown in Fig. 5, in which data from three experiments are given (open symbols). A four-parameter logistic curve was fitted to the average values for the data points, with the constraint that the curve pass through the average values for Rmin and Rmax. The regression curve produced values for the steepness parameter n, representing the number of maximally cooperative binding sites for Ca2+ on fura 2 needed to construct the curve, and the position parameter Kd · beta , as shown in Fig. 5. The value for beta  was obtained as described in METHODS (see Calibration). The value for n is close to 1, as would be expected for noncooperative binding to a single site. These values are all in reasonable agreement with those reported previously for an in vitro calibration of fura 2 (17).


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Fig. 5.   In vitro and in vivo calibration of fura 2 fluorescence ratio (R) signal. In each case, results are plotted from 3 separate experiments: open symbols, in vitro; filled symbols, in vivo. Temperature was 30°C. A logit equation (SigmaPlot 4.0) was used to fit a curve to each group of points. Basic equation relating Ca2+ concentration ([Ca2+]) to R is [Ca2+] = Kd · beta  · [(R - Rmin)/(Rmax - R)], where Kd is the dissociation constant, beta  is the ratio of Ca2+-free to Ca2+-bound fluorescence at 380 nm, and Rmin and Rmax are the minimum and maximum fluorescence ratios, respectively. For calibration, this equation must be rewritten to express R as a function of [Ca2+]. Thus R = [([Ca2+] · Rmax) + (Kd · beta  · Rmin)]/(Kd · beta  + [Ca2+]), which can be written as R = (Rmax - Rmin)/[1 + (Kd · beta /[Ca2+]n)] + Rmin, where n is the steepness parameter. Four parameters must be found: Rmax, Rmin, Kd · beta , and n, which is ordinarily near 1, because it is assumed that only 1 binding site for Ca2+ is present on fura 2. In the regression, parameters Rmax and Rmin were constrained to their respective average values, whereas n and Kd · beta  were not. Note that a parameter Kd · beta  remains; therefore, to obtain Kd it is necessary to divide by beta . Procedure to obtain beta  is described in METHODS. Av.beta means the averaged beta  value from 3 different experiments.

For in vivo calibration of fura 2, attention was particularly directed toward avoiding significant changes in autofluorescence. The main component of autofluorescence in rat trabeculae is the result of NADH, even under optimal optical conditions for fura 2 fluorescence measurement, and it varies substantially, particularly when metabolic inhibition occurs, as shown in Fig. 2. Therefore, we chose to keep the normal metabolic pathway uninterrupted during an in vivo calibration. The rationale behind this is that a constant internal concentration of ATP ([ATP]i) should result in a constant [NADH]i when the normal metabolic pathway is intact. We used three inhibitors of ATPases simultaneously in our calibration solutions: BDM (also blocking contracture), TBQ, and strophanthidin, which block, respectively, actomyosin-ATPase, SR Ca2+-ATPase, and Na+-K+-ATPase. Under these conditions, the effects of the Rmin and Rmax solutions were examined on excitation wavelength scans of tissue autofluorescence in rat trabeculae without loading fura 2 at 30°C (corrected by subtracting out the background fluorescence). In both Rmin and Rmax solution at 380 nm, the treated curves produced an autofluorescence ~8% greater than control, whereas at 345 nm the treated autofluorescence was ~22% greater than the control (data not shown). A difference spectrum (treated control) showed a wavelength distribution very similar to that of NADH. A likely explanation is that both the solutions contain potent inhibitors of ATPases, thus blocking ATP usage with consequent rise in NADH. These small changes in autofluorescence caused increases in the estimates of Rmin and Rmax by ~1 and 2%, respectively, when dye fluorescence was 10 times that of the combined background fluorescence and autofluorescence. This insignificant change in the R values allowed us to calibrate fura 2 in vivo without correction of autofluorescence increase caused by calibration solutions.

The results of in vivo calibration are also shown in Fig. 5, in which the same nonlinear regression function, with the same constraints for Rmax and Rmin, was used to generate a curve to fit the averaged data points from three trabeculae (solid symbols). A comparison of the in vitro and in vivo results shown in Fig. 5 reveals quite different results for Rmax (20.8 vs. 7.97), beta (13.4 vs. 6.9), and Kd (201 vs. 678 nM). These results are not in agreement with previously published ones for this preparation at 22°C (3). Intracellularly, dye properties could change as a result of binding to other myoplasmic constituents (5), and, also, myoplasmic viscosity could be responsible for changing the properties of fura 2 (27). It has been reported that the in vivo properties of indo 1 in myocytes change substantially, yielding an apparent Kd several times higher than that obtained in vitro (4). Also, similar in vivo shifts of the spectral as well as the Ca2+-binding properties of other fluorescent dyes have been reported (18, 25), as indicated by the much smaller in vivo Rmax. Whatever the explanation for the in vivo spectral shifts and changes in Ca2+-binding properties of fura 2, the in vivo fitted calibration curve shown in Fig. 5 makes it possible to transform our results for the R signal into [Ca2+]i using the original equation (17) with parameter values appropriate for the intracellular environment.

Because both Rmax and Kd were altered in the myoplasm, the [Ca2+]i inferred from R during a twitch with the parameters from in vitro or in vivo calibration must be very different. The results are shown in Fig. 6 for a typical R signal obtained in these series of experiments. The appropriate values for Rmax, Rmin, and Kd · beta  were taken from the data shown in Fig. 5. Note the very substantial differences between the diastolic and peak systolic values for [Ca2+]i, depending on which calibration is used. These results emphasize the necessity of doing a complete in vivo calibration curve when using fura 2 if reliable results concerning the true values for [Ca2+]i are required. This requires not only obtaining values for Rmax and Rmin but also determining Kd · beta  to make it possible to express [Ca2+]i with reasonable confidence over the complete range of R from Rmin to Rmax. Comparable results have been presented in which the in vivo Kd for indo 1 in rabbit myocytes increased by about a factor of 3.4 (4), which is similar to that found by us for fura 2. 


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Fig. 6.   R signal of a twitch and its transformation into intracellular concentration of Ca2+ ([Ca2+]i), using parameters from in vitro and in vivo calibrations. A: a typical R signal is shown (average of 18 interleaved sweeps, with 9 at 345 nm and 9 at 380 nm) as a function of time at 30°C and pacing of 0.5 Hz. B and C: transformations into [Ca2+]i are shown using in vitro and in vivo calibration parameters, respectively. Values for Rmax, Rmin, and Kd · beta  used in the 2 transformations were taken from data shown in Fig. 5. Transforming equation was [Ca2+]i = Kd · beta  · [(R - Rmin)/(Rmax - R)]. Note substantial differences in [Ca2+]i as a function of time, depending on which calibration is used.

As mentioned above, using our new calibration method, we were able to reduce the changes of autofluorescence during the calibration to levels that were insignificant when fura 2 fluorescence was 10 times higher than the autofluorescence. It is not desirable to load fura 2 to such a high degree to measure [Ca2+]i transients, as it may significantly buffer Ca2+ and reduce the peak value of R (5). We ordinarily load fura 2 to an extent that the total fluorescence after loading is about three times higher than the initial autofluorescence. Because high pacing frequency, halothane, and BDM cause significant autofluorescence changes, proper correction of recorded fluorescence for the autofluorescence may result in a very different R value from that found by assuming constant autofluorescence. Figure 7 shows the difference in the R signals (A) and the corresponding [Ca2+]i traces (B) with and without taking account of the autofluorescence change resulting from the high pacing frequency. If we assume that autofluorescence does not change after the pacing frequency is switched from 0.5 Hz to 5 Hz and simply subtract the autofluorescence at 0.5 Hz from the raw fluorescence excited at 345 nm (F345) and at 380 nm (F380) at 5 Hz, the results are an improperly corrected R trace and corresponding incorrect [Ca2+]i. Properly correcting the autofluorescence changes during the 8 s of 5-Hz pacing revealed the differences, compared with those that were improperly corrected, in the R and corresponding [Ca2+]i traces. The change in [Ca2+]i between the properly and improperly corrected traces is shown in the difference plot at the bottom of Fig. 7B. At the initiation of the 5-Hz pacing, the difference was small and gradually become more prominent. Then, 4 s after the last stimulus at 5 Hz, the difference between the corrected and uncorrected traces disappeared. Figure 7C shows the difference between the uncorrected and corrected [Ca2+]i, using an expanded time base to cover the succeeding 4 s after the last stimulus at 5 Hz.


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Fig. 7.   R signal and [Ca2+]i of a trabecula with proper correction for change in autofluorescence and improper correction assuming constant autofluorescence. Autofluorescence at 345 nm (F345) and at 380 nm (F380) was recorded using 2 pacing protocols (1 and 2), and photon-counting mode was used with a data collection rate of 200/s. In protocol 1, preparation was continuously paced at 0.5 Hz. In protocol 2, preparation was paced at 0.5 Hz for 20 s, with an intervening period of 8 s at 5 Hz. F345 and F380 were averages of 9 interleaved measurements, using both protocols. After loading of fura 2, raw F345 and F380 were collected using protocol 2 and were also averages of 9 measurements. A: R signal (thin line) was formed by division of F345 by F380 for both 0.5- and 5-Hz pacing after subtraction of autofluorescence obtained using protocol 1 (improper correction). Also in A, with use of protocol 2, R signal (thick line) is shown corrected by subtraction of autofluorescence change caused by switching pacing frequency from 0.5 to 5 Hz (proper correction). B: corresponding [Ca2+]i of R signals in A were obtained by transformation, using parameters obtained from in vivo calibration. Thin line shows improperly corrected trace, whereas thick line shows properly corrected trace. Difference (proper - improper) trace at the bottom of B shows more clearly the change of [Ca2+]i during 5-Hz pacing obscured by associated change in autofluorescence. C: data from B are plotted using an expanded time base from 12 to 16 s and an ordinate scale from 0 to 1, and elevation in [Ca2+]i (proper correction, thick line) returned to baseline level associated with 0.5-Hz pacing (improper correction, thin line) within ~4 s after last stimulus at 5-Hz pacing. Note in this trabecula alternating variation in R signal amplitude associated with pulsus alternans in the twitch response (not shown).

    DISCUSSION
Top
Abstract
Introduction
Methods
Results
Discussion
References

The results presented here emphasize that the strong autofluorescence of cardiac muscle can be excited by incident light in the 300- to 450-nm range (emission at 510 nm) in an apparatus optimized for use of fura 2. As a result mainly of the prolific work by Chance and Thorell (11) and their colleages, it is now generally appreciated that the cellular basis for this autofluorescence resides in NADH in mitochondria, and it is confirmed in this work that changes in autofluorescence correlate with expected changes in NADH level. Our results also indicate that autofluorescence in heart is not constant and that it can vary considerably under various conditions, e.g., various pacing protocols. Because the excitation spectra of fura 2 and the autofluoresecence overlap with each other, the change in autofluorescence seriously affects the use of fura 2 in the heart. To use fura 2, as well as other dyes that have similar excitation and emission wavelengths in the heart, it is necessary to first determine how NADH varies under all conditions encountered in the experimental protocol.

One way to minimize the effect of the autofluorescence change on obtaining valid estimations of R is to make the autofluorescence change relatively smaller by loading fura 2 fluorescence to a level well above the initial autofluorescence level. However, this is limited by the Ca2+-buffering capacity of fura 2, because a high concentration of fura 2 substantially decreases [Ca2+]i and twitch force. Typically, in our work, the fluorescence intensity after loading fura 2 is about three times that of the initial autofluorescence. Another way to circumvent the problem is to choose proper experimental conditions in which the autofluorescence is kept relatively constant. Because [ATP]i is coupled to [NADH]i, keeping the [ATP]i constant would minimize the change in autofluorescence level. We found in this study that a change in pacing frequency from 0 to 0.5 Hz or vice versa caused little change in autofluorescence level (data not shown). One explanation for this is that, at a low pacing rate (<= 0.5 Hz), the ATP consumption rate is low and [ATP]i is relatively constant. Under the conditions of 30°C, 2 mM [Ca2+]o, and pacing rate of <= 0.5 Hz, together with loading dye fluorescence three times that of the initial autofluorescence, reliable R values could be obtained without correction for a change in autofluorescence.

Once a reliable R value is obtained, a correct transformation from R to [Ca2+]i is critical for an accurate estimation of [Ca2+]i, and a calibration is crucial to do this. It has been reported that the Kd for fura 2 is near 224 nM, at least for in vitro aqueous buffer media (13). If this is the case, it can be shown that, even with [Ca2+]i near 1 µM, substantial nonlinearity of response would occur. This makes it necessary to calibrate R using well-defined buffer media that is usually EGTA based in which the [Ca2+] is known and constant (6). However, it is now known that the calibration ideally should be carried out under in vivo conditions (32). This is difficult in cardiac muscle because the tissue tends to contract strongly, particularly in the solution used to determine Rmax, and also because powerful cellular pumps exist, e.g., the Na+/Ca2+ exchanger, that act to resist any attempt to change [Ca2+]i (4). A common method in the field is the metabolic inhibition technique (23). This technique has been applied to rat trabeculae like those used here about which it was reported that metabolic inhibition produced only small changes in autofluorescence (3). This is surprising because under these conditions production of ATP should be profoundly inhibited, with a concomitant change in the NADH level either as a result of the cyanide or iodoacetate treatment, which would produce opposite effects on the NADH level. As shown in our Fig. 2 and by others (8), cyanide raises the level of NADH, whereas FCCP decreases it. It has also been reported that autofluorescence levels measured before loading with fura 2 and again at the end of an experiment by quenching fura 2 fluorescence with Mn2+ differed from each other by <2% (2). Others have also used the metabolic inhibition technique to calibrate fura 2 in vivo without any apparent comment about the influence on autofluorescence (16). Usually, no attempt is made to measure beta , so it was only possible to give the apparent Kd, i.e., Kd · beta . As has been pointed out, Kd, or the apparent Kd, i.e., Kd · beta , enters into the equation relating [Ca2+]i to R as a linear scale factor (4).

In our view, the conditions necessary to produce metabolic inhibition are extreme. Cyanide and FCCP cause the oxidation-reduction state to change from one extreme level to another. This greatly alters the autofluorescence and, in turn, could lead to incorrect R values. If it is assumed that the metabolic inhibition technique used for calibration (23) does decrease both F345 and F380 of the autofluorescence to the same degree as FCCP, as shown in Fig. 2A, and that the total fluorescence after loading fura 2 is only three times the initial autofluorescence, the Rmin and Rmax values would become 0.44 and 23.8, respectively, which by chance are close to those of the in vitro calibration, instead of 0.72 and 8.2, respectively, obtained under our conditions. This great disparity in Rmin and Rmax values will produce a large difference in the transformation of R into [Ca2+]i. These problems were avoided in our approach by greatly decreasing ATP consumption, which kept the NADH level near constant. This was done by blocking actomyosin ATPase (BDM), SR Ca2+ pump ATPase (TBQ), and membrane Na+-K+-ATPase (strophanthidin). Indeed, our results show that the effect of these calibrating solutions on autofluorescence is minimal, and this, together with dye fluorescence in great excess over autofluorescence, makes it very likely that only dye fluorescence enters into the calibration protocol (see Calibration in METHODS). A consequence of our in vivo calibration protocol is that the value after transforming R for peak systolic [Ca2+]i is necessarily higher than generally reported, as shown in Fig. 6. Obviously, it is important to know whether peak systolic [Ca2+]i is ~2 µM as reported here for 30°C and 2 mM [Ca2+]o, or whether it is nearer 600-700 nM, as previously reported for 22°C and 1 mM [Ca2+]o (3).

With the correct choice of experimental conditions together with a reliable calibration, [Ca2+]i could be estimated with confidence. However, in some conditions, such as pacing at high frequency, autofluorescence change cannot be avoided. Under this condition, muscle work is greatly increased, which causes an increase in ATP consumption, thus leading to a decrease in NADH level as shown in Fig. 3 and by others (8, 9). A similar effect could occur when muscle work is rapidly reduced by other agents, such as halothane and BDM. However, effects of these two agents on the autofluorescence level of cardiac muscle depend on pacing frequency, as shown in Fig. 4. Both halothane and BDM cause substantial increases in autofluorescence only when paced at high frequency (e.g., 5 Hz), at which the autofluorescence increases to approach the level at 0.5-Hz pacing, as shown in Fig. 4. This suggests that both halothane and BDM increase the autofluorescence, or NADH level, at high pacing rates simply by reducing muscle work (and ATP consumption). At a low pacing rate, e.g., 0.5 Hz, ATP consumption is low and the NADH (and autofluorescence) is already at a high level. Here, treatment with halothane or BDM to decrease muscle work does not cause an appreciable further decrease in ATP consumption, and the NADH level does not rise significantly.

If the change in autofluorescence level is large enough to interfere with obtaining an accurate R value, an appropriate correction for the autofluorescence change is necessary to use fura 2 to report [Ca2+]i. It is clearly shown in Fig. 7 that the improperly corrected trace erroneously indicates that the [Ca2+]i rapidly returned to the diastolic level at 12.5 s, although the actual [Ca2+]i is still ~250 nM higher at that time. In addition, the proper correction also affects the interpretation of the kinetics of the [Ca2+]i fall. In Fig. 7, the fall in [Ca2+]i, indicated by the properly corrected trace, could be well fit with a double exponential curve with decay rates of 11.8 counts/s and 0.3 counts/s. However, the improperly corrected trace, unreliable owing to concurrent autofluorescence changes, was well fit using a single exponential with a decay rate of 9.5 counts/s. This problem is even more serious when the pacing frequency is switched from high (5 Hz) to no stimulation (0 Hz) or vice versa, as shown in Fig. 3. Termination of pacing at high frequency results in a rise and overshoot to a higher value in autofluorescence (9). In summary, autofluorescence changes in the heart interfere with fura 2 reporting of [Ca2+]i. Interventions causing a large change in autofluorescence, if not properly corrected, will result in an inaccurate estimate of [Ca2+]i both transiently and in the steady state.

    ACKNOWLEDGEMENTS

This work was supported by National Institutes of General Medical Sciences Grant GM-48078 (F. J. Julian).

    FOOTNOTES

Address for reprint requests: Y. Jiang, Dept. of Anesthesia Research Laboratories, Brigham and Women's Hospital, 75 Francis St., Boston, MA 02115.

Received 7 July 1997; accepted in final form 22 August 1997.

    REFERENCES
Top
Abstract
Introduction
Methods
Results
Discussion
References

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