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. 2015 Mar 13;593(Pt 8):1829–1840. doi: 10.1113/jphysiol.2014.286153

Rapid changes in NADH and flavin autofluorescence in rat cardiac trabeculae reveal large mitochondrial complex II reserve capacity

Rob C I Wüst 1,✉, Michiel Helmes 1,2, Ger J M Stienen 1,3
PMCID: PMC4405745  PMID: 25640645

Abstract

The functional properties of cardiac mitochondria in intact preparations have been mainly studied by measurements of nicotinamide adenine dinucleotide (NADH) autofluorescence, which reflects mitochondrial complex I function. To assess complex II function, we extended this method by measuring flavin adenine dinucleotide (FAD)-related autofluorescence in electrically stimulated cardiac trabeculae isolated from the right ventricle from the rat at 27°C. NADH and FAD autofluorescence and tension responses were measured when stimulation frequency was increased from 0.5 Hz to 1, 2 or 3 Hz for 3 min, and thereafter decreased to 0.5 Hz. Maximal complex I and complex II activity in vitro were determined in saponin-permeabilized right ventricular tissue by respirometry. NADH responses upon an increase in stimulation frequency showed a rapid decline, followed by a slow recovery towards the initial level. FAD responses followed a similar time course, but in the opposite direction. The amplitudes of early rapid changes in the NADH and FAD concentration correlated well with the change in tension time integral per second (R2 = 0.833 and 0.660 for NADH and FAD, respectively), but with different slopes for the up and down transient. Maximal velocity of the increase in FAD concentration (16 ± 4 μm s−1), measured upon an increase in stimulation frequency from 0.5 to 3 Hz was considerably smaller than that of the decrease in NADH (78 ± 13 μm s−1). The respiration measurements indicated that the maximal velocity of NADH utilization (143 ± 14 μm s−1) was 2 times smaller than that of FADH2 (291 ± 19 μm s−1). This indicates that in cardiac mitochondria considerable complex II activity reserve is present.

Key points

  • A photometry-based technique was developed to measure nicotinamide adenine dinucleotide (NADH) and flavin adenine dinucleotide (FAD) autofluorescence and contractile properties simultaneously in intact rat trabeculae at a high time resolution. This provides insight into the function of mitochondrial complex I and II.

  • Maximal complex I and complex II activities were determined in saponin-permeabilized right ventricular tissue by respirometry.

  • In trabeculae, complex II function was considerably smaller than the maximal complex II activity, suggesting large complex II reserve capacity.

  • Up–down asymmetry in NADH and FAD kinetics suggests a complex interaction between mitochondrial and contractile function.

  • These data show that simultaneous measurement of contractile properties and NADH and FAD kinetics in cardiac trabeculae provides a mean to study the differences in complex I and II function in intact preparations in health and disease.

Introduction

The heart has the ability to increase its mechanical output during exercise and stress several fold within seconds. This is accompanied by a rapid increase in the rate of oxidative phosphorylation in the mitochondria (for a review see Balaban, 2009).

Mitochondrial complex I and II serve as the major sites of electron input into the electron transport system through nicotinamide adenine dinucleotide (NADH)- and reduced flavin adenine dinucleotide (FADH2)-coupled electron transfer. Since the pioneering work of Chance and colleagues (Chance et al. 1967, 1972; Scholz et al. 1969), autofluorescence of reduced NADH has been used as a measure of the activity of complex I of the electron transport system, allowing inferences to be made about the control of oxidative phosphorylation (Brandes & Bers, 1996, 1999, 2002; Brandes et al. 1998; Bose et al. 2003; Hogan et al. 2005; Wengrowski et al. 2014). The studies of Bers and coworkers indicated that the NADH concentration decreases rapidly during the transition from low to high stimulation frequency and stabilizes to a new steady-state level in rat cardiac trabeculae (Brandes & Bers, 1996), indicative for rapid oxidation of NADH in complex I at the start of the transition from low to high stimulation frequency.

Electrons from NADH into mitochondrial complex I and from FADH2 into complex II are transferred into complex III by convergent input into the Q10 complex (Gnaiger, 2009). Recently, it has been shown that electron flux is determined by dynamic supercomplexes whose organization depends on the available substrates (Lapuente-Brun et al. 2013). Little is known about the kinetics of complex II function in intact preparations. FAD is the oxidized form of FADH2, utilized in complex II and thus reflects mitochondrial complex II activity. The flavin autofluorescence spectrum contains three main components with maxima at 500, 530 and 560 nm. The flavins bound to α-lipoamide dehydrogenase and electron transferring flavoprotein contribute to the 500 nm component and the 530 nm component originates from free FAD (Chorvat et al. 2005; Sedlic et al. 2010), and is predominantly due to mitochondrial flavoproteins (Scholz et al. 1969). This suggests that fluorescence measured in the range from 500 to 540 nm can be used to assess complex II function. Therefore, we extended this photometry-based technique to measure NADH and FAD fluorescence and contractile properties simultaneously in intact rat trabeculae at a much higher time resolution than achieved previously (e.g. Aldakkak et al. 2008). Using this method, we investigated whether FAD transients upon changes in stimulation frequency show opposite responses compared to NADH.

Complex I and II activities are probably intensity-dependent because of Ca2+-activated mitochondrial respiration (Brandes & Bers, 1996, 1999; Cortassa et al. 2003; Glancy & Balaban, 2012). Recent evidence in skeletal (Scorzeto et al. 2013) and cardiac muscle (Drago et al. 2012) has provided evidence that rapid asymmetric increases and decreases in mitochondrial free Ca2+ concentration cause gradual Ca2+ accumulation upon an increase in stimulation frequency. This mitochondrial Ca2+ accumulation is suggested to cause activation of mitochondrial transporters and/or enzyme activities, effectively resulting in an increase in oxidative phosphorylation (Balaban, 2009; Glancy & Balaban, 2012). A consequence of mitochondrial Ca2+ accumulation is that the NADH and FAD responses probably show up–down asymmetry upon an increase and decrease in contractile output (cf. Wüst et al. 2013). Therefore, we also studied the changes in NADH and FAD concentration upon an increase as well as a decrease in contractile output.

Parallel experiments in permeabilized cardiac tissue were performed to determine the maximal complex I- and complex II-coupled respiration. These measurements allowed us to compare the measurements of net NADH and FADH2 utilization in intact preparations with the maximal complex I and II activities.

Methods

Animals and ethical approval

All protocols were in accordance to ARRIVE guidelines and were approved by the Animal Experimental Welfare Committee of the VU University Medical Center Amsterdam, where all experiments were performed.

Surgical preparation

The isolation procedure of the cardiac trabeculae was as described previously (Lamberts et al. 2007). In short, male Wistar rats (∼300 g, n = 6) were anaesthetized with isoflurane. The heart was rapidly removed and retrogradely (Langendorff) perfused with a modified Krebs–Henseleit (tyrode) solution, consisting of (in mm): NaCl (118), KCl (4.5), CaCl2 (0.5), NaH2PO4 (0.33), MgCl2 (1), NaHCO3 (25) and glucose (10), gassed with 95% O2/5% CO2 at pH 7.45, and supplemented with 2,3-butanedione monoxime (20) to inhibit cross-bridge cycling. A thin trabecula (diameter <200 μm) from the right ventricle was dissected and mounted in an experimental chamber between a force transducer and a micromanipulator. The trabecula was constantly superfused with tyrode solution (without 2,3-butanedione monoxime) to guarantee adequate oxygen supply. The trabecula was supramaximally stimulated (∼1.5× threshold voltage; pulse width 1 ms) using a field stimulator (IonOptix Myopacer EP; IonOptix LLC, Milton, MA, USA). The force transducer signal (HJK, Merching, Germany) was filtered with a 100 Hz low-pass filter. Optimal length of the preparation was determined while stimulating the trabecula at 0.5 Hz and stretching the preparation until developed tension was maximal. At optimal length, the diameters were measured in two perpendicular directions and the cross-sectional area was calculated assuming an elliptical cross-section. Temperature was kept at 27°C to maintain stability of the preparation during the experiments.

Experimental setup

The experimental chamber with the trabecula was placed on the stage of an inverted microscope (AE31; Motic, Richmond, BC, Canada) with a 10× objective (UPLSAPO 10×; Olympus; Zoeterwoude, the Netherlands) that is designed for optimal transmission of light at 340 nm. Figure 1 shows an overview of the setup.

Figure 1. Schematic overview of experimental setup.

Figure 1

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Thin cardiac trabecula mounted between a force transducer and a micromanipulator to adjust its length was superfused with tyrode solution and electrically stimulated at different stimulation frequencies. Using a 10× objective and a series of dichroic mirrors, filters and two PMTs, NADH and FAD autofluorescence as well as an image of the trabecula were recorded. PMT, photomultiplier tubes.

A 75 W Xenon lamp (USHIO, Tokyo, Japan), set to 65 W, was used for illumination and a HyperSwitch (IonOptix; LLC, Milton, MA, USA) was used for rapid wavelength switching. The central part of the trabecula was illuminated at 340 nm to assess autofluorescence of NADH, immediately followed by illumination at 480 nm for measurement of FAD autofluorescence. A rectangular mask was used to select light emitted from the central region of the trabecula. Filters and dichroic mirrors were purchased from Chroma Technology (Brattleboro, VT, USA) unless indicated otherwise. Emitted light was passed through a long-pass dichroic mirror with a cut-off frequency of 355 nm and a band stop filter at about 488 nm (Z355/488RPC; Semrock, Rochester, NY, USA). A dichroic mirror (585 nm long pass; 585DCXR) was used to separate the fluorescence light from the red filtered light used for transmission illumination of the trabecula. An image of the trabecula was displayed on a CCD camera (MyoCam-S, IonOptix). Emitted light with a wavelength of 455 nm was selected with a dichroic mirror (T495LPXR) and filter (ZET455/10X) and guided to a photomultiplier tube (PMT; H7360-02MOD; Hamamatsu Photonics, Hamamatsu, Japan) for the detection of NADH autofluorescence. In cardiac tissue, the NADH fluorescence signal primarily arises from the mitochondrial pool (Eng et al. 1989) and the contribution of NADPH is negligible (Klingenberg & Slenczka, 1959).

FAD-related autofluorescence in the range from 500 and 540 nm was detected with a second PMT using a bandpass filter (Z520/20X). As mentioned in the Introduction, this signal originates from three different flavin compounds. The autofluorescence in the range from 500 to 540 nm was most affected by cyanide and mitochondrial uncoupling in cardiomyocytes and therefore closely resembles the emission reported for free oxidized FAD (Scholz et al. 1969; Chorvat et al. 2005). Therefore we refer to this signal as FAD autofluorescence. To minimize movement artefacts and bleaching, autofluorescence of NADH and FAD were detected during two successive periods of 10 ms each at 1 s intervals, timed during the final diastolic phase of the contraction (open circles, Fig. 2).

Figure 2. Typical example of NADH and FAD autofluorescence at high (250 Hz) illumination frequency upon a change in stimulation frequency from 0.5 to 3 Hz.

Figure 2

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Tension recording (A), NADH autofluorescence (B) and FAD autofluorescence (C) during glucose superfusion. During the tension transient movement, artefacts are visible. The fluorescence recordings shown in Figs 4 and 5 were obtained at 1 Hz illumination frequency (duration of 10 ms) at the time points indicated by the open circles. Upon rapid changes in stimulation frequency, at t = 0 (s), contractile properties and diastolic NADH and FAD autofluorescence change within seconds, but stabilize afterwards (see Fig. 3). Similar changes but in opposite direction were observed during the transient responses to back to a stimulation frequency of 0.5 Hz (down transient).

The concentration dependence of the NADH and FAD signals and cross-over between them were assessed using thin capillaries filled with known concentrations of NADH and FAD. Up to the maximum concentrations expected for NADH and FAD, approximately 2.5 and 0.3 mm, respectively, intensity varied in proportion to concentration (NADH: R2 = 0.996; FAD: R2 = 0.988). Cross-over from the FAD into the NADH signal was negligible, while the contribution of NADH in the FAD signal was <5%. Stray light from the HyperSwitch at 340 nm resulted in slight contamination (1.3%) during illumination at 480 nm, whereas the converse (480 nm stray light during 340 nm illumination) was less than 0.2%. To stay in the linear range of the photon counting domain of the PMT, neutral density filters were placed into the light path to keep the fluorescence intensity values <7000.

A photo-spectrometer (QE65Pro; Ocean Optics, Dunedin, FL, USA) was used to test the overall emission spectra of the trabeculae at the two excitation wavelengths used. No additional peaks were observed in the spectrum between 400 and 600 nm.

Experimental protocol

After switching to tyrode solution with 1.0 mm CaCl2 trabeculae were allowed to stabilize for >1 h. The experimental protocol was started by recording force and NADH and FAD autofluorescence signals at a stimulation frequency of 0.5 Hz for 3 min. Thereafter, the trabecula was stimulated at a higher stimulation frequency of 1, 2 and 3 Hz for at least 3 min to study the tension–frequency relationship.

After a stabilization period of at least 30 min at 0.5 Hz stimulation, the trabecula was stimulated at 1, 2 or 3 Hz for 3 min, followed by a recovery period at 0.5 Hz of at least 30 min. Stimulation sequences at 1, 2 and 3 Hz were selected at random.

Calculations

Maximal (peak) tension, i.e. peak force divided by the cross-sectional area of the preparation (Tpeak in mN mm−2), was determined for each contraction during the experimental runs. The steady-state Tpeak reached was used to construct the tension–frequency relationship. The tension time integral per second (TTI/s, in mN mm−2) was determined throughout the experimental run and is equal to the time-averaged developed tension. It is a good indicator of total mechanical output and the associated energy utilization.

Background values for the NADH and FAD intensity were subtracted and all values during each experimental run were normalized to the initial value (100%). A very marginal rundown of NADH and FAD autofluorescence over the experimental run of ∼15 min was accounted for by correcting the slope of the decline in autofluorescence. Separate experiments (n = 5) in which 5 mm cyanide and 4 μm carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) were used to maximally reduce and oxidize the total pyridine nucleotide and the total flavin pool (Brandes & Bers, 1996), showed that the pyridine nucleotide pool in these trabeculae was 63 ± 6% reduced at 0.5 Hz. Similarly, the flavin pool was 57 ± 11% oxidized at 0.5 Hz.

The maximal velocities of the early changes in NADH and FAD autofluorescence upon an increase or decrease in stimulation frequency were determined from the differentiated signal (in % s−1). To express the maximal velocities in μm s−1, the total pyridine nucleotide concentration inside the mitochondrial matrix was assumed 3.4 mm (Joubert et al. 2004). This would result in a NADH concentration of 2.14 mm. To estimate the total flavin concentration, we determined the in vitro relations between autofluorescence intensity and the NADH and FAD concentration in small (20 and 50 μl) droplets as well as the average ratio of NADH to FAD autofluorescence intensity in the trabeculae (n = 6) during 0.5 Hz stimulation at the start of the measurements (1.05 ± 0.07). Using the in vitro calibrations, this value would correspond with a FAD concentration of 0.30 mm and the total flavin concentration would amount to 0.53 mm.

Mitochondrial respirometry

Mitochondrial respiration was determined in permeabilized cardiac tissue (n = 6) as described previously (Wüst et al. 2012) with small modifications. Briefly, small right ventricular tissue samples (∼3 mg wet weight) were permeabilized with 50 μg ml−1 saponin for 30 min at 4°C in preservation solution, consisting of (in mm) CaK2EGTA (2.8), K2EGTA (7.2), ATP (5.8), MgCl2 (6.6), taurine (20), phosphocreatine (15), imidazole (20), DTT (0.5) and MES (50); pH 7.1, adjusted with potassium hydroxide. Tissue was subsequently washed in mitochondrial respiration solution (MiR05), containing EGTA (0.5), MgCl2 (3), potassium lactobionate (60), taurine (20), KH2PO4 (10) Hepes (20), sucrose (110) and 1 g l−1 fatty acid free bovine serum albumin (pH 7.1), quickly blotted dry, weighed and transferred to a high-resolution respirometer (Oxygraph-2k; Oroboros Instruments, Innsbruck, Austria).

The oxygen concentration was maintained above 300 μm throughout the measurements to avoid limitations in oxygen supply. Maximal complex I-coupled respiration was measured in MiR05 after addition of 10 mm sodium glutamate, 2 mm sodium malate and 5 mm sodium pyruvate and 2.5 mm ADP. The outer mitochondrial membrane integrity was tested by the addition of 10 μm cytochrome c. Maximal oxidative phosphorylation, with simultaneous input of electrons through complex I and II, was measured after addition of 10 mm succinate. Subsequently, complex II-coupled respiration was measured after blocking complex I by adding 0.5 μm rotenone. Residual oxygen consumption, measured in the presence of 2.5 μm antimycin A, was subtracted from all values. Measurements were performed at 37°C. Two tissue samples from the same heart were measured simultaneously and the results were subsequently averaged.

To allow a comparison with the measurements in intact trabeculae, the temperature coefficients (Q10) for mitochondrial respiration were studied in a separate group of samples (n = 4) by adjusting the temperature during the course of the experiment to 37 and 27°C. The temperature-dependent instrumental background and calibration were taken into account in the evaluation of the results.

Values obtained were expressed per mg wet weight and converted from pmol O2 s−1 mg−1 to μm s−1 NADH or FAD, assuming mitochondrial volume to be 36% of the total volume (Cortassa et al. 2006).

Statistical analysis

Comparisons were made using repeated measures ANOVA (with stimulation frequency and/or an increase/decrease in stimulation frequency as within factors, followed by Bonferroni post-hoc tests) or paired t tests where appropriate. The level of significance was set at P < 0.05. Statistical analyses were performed using SPSS 20.0 (IBM SPSS Statistics, Armonk, NY, USA). All data are presented as means ± SEM.

Results

High time resolution experiments

NADH and FAD autofluorescence measurements with a repetition frequency of 250 Hz and illumination duration of 2 ms for NADH and FAD were performed to check whether the recordings were affected by bleaching, intra-beat changes and/or movement artefacts of the contracting trabecula. Figure 2 depicts a typical example of a recording during a change in stimulation frequency from 0.5 to 3 Hz. Tension followed the pattern described previously (Schouten, 1990). The NADH and FAD autofluorescence signals increased by 5–10% during tension development, as expected from shortening of the central part of the trabecula at the expense of the end regions. No other systematic intra-beat changes were observed. Bleaching was most pronounced in the NADH signal and amounted to 3.6 ± 0.3% min−1 (n = 3). To minimize the influence of movement artefacts and bleaching, the NADH and FAD signals in subsequent experiments were recorded at 1 s intervals and illumination duration was set at 10 ms for both NADH and FAD just before the stimulation pulse. Bleaching under these conditions was very small as total illumination duration amounted to only 2% of that during the 250 Hz recordings.

Contractile properties

Figure 3A shows the typical steady-state tension responses observed at 0.5, 1, 2 and 3 Hz stimulation frequency. Increasing the stimulation frequency resulted in a larger peak tension, but also shortened the contractions: both the time to peak and total contraction time were shorter at higher stimulation frequencies. Accordingly, the relationship between amplitude of the tension transient and stimulation frequency was positive (P = 0.001; Fig. 3B). The TTI/s increased linearly with stimulation frequency (Fig. 3C). A detailed overview of the contractile properties at different stimulation frequencies is presented in Table1.

Figure 3. Contractile properties at different stimulation frequencies.

Figure 3

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A, tension responses at stimulation frequencies of 0.5, 1, 2 and 3 Hz. Both amplitude and duration of the tension transients were different across stimulation frequencies (see also Table1). B, positive tension–frequency relationship was observed, C, tension time integral per second (TTI/s) was linearly related to stimulation frequency. Means ± SEM (n = 6). *P < 0.05 compared 0.5 Hz.

Table 1.

Contractile properties of rat trabeculae

0.5 Hz 1 Hz 2 Hz 3 Hz
Tpeak 21.8 ± 2.5 23.2 ± 1.4 27.2 ± 4.0* 30.1 ± 3.9*
MRC 12.9 ± 0.2 13.8 ± 0.3* 15.1 ± 0.2* 16.7 ± 0.4*
MRR 7.9 ± 0.6 8.3 ± 0.5 9.4 ± 0.8* 10.7 ± 1.0*
TTP 134 ± 3 124 ± 3* 109 ± 2* 99 ± 3*
TCT 402 ± 26 375 ± 25* 306 ± 20* 253 ± 13*
TTI/s 1.9 ± 0.3 3.0 ± 0.6* 5.8 ± 0.9* 10.4 ± 1.4*
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MRC, maximal rate of contraction, normalized for Tpeak (s−1); MRR, maximal rate of relaxation (s−1); TCT, total contraction time (ms); Tpeak, maximal tension (mN mm−2); TTI/s, tension time integral per s (mN mm−2), which is equal to average tension; TTP, time to peak (ms). Values are means ± SEM.

*Significantly different from 0.5 Hz (P < 0.05).

NADH and FAD autofluorescence

Baseline values

The NADH autofluorescence baseline values increased by approximately 20% during the initial stabilization phase (1 h) at 0.5 Hz and during the actual measurements it declined gradually to 80% of the initial value. The FAD autofluorescence baseline signal was very stable.

NADH and FAD autofluorescence responses

Overall, the NADH and FAD responses upon a change in stimulation frequency changed in opposite directions (Fig. 4). The NADH and FAD responses consist of an early rapid response followed by a slow recovery phase. The early rapid response upon an increase in stimulation frequency coincides with the increase in contractile output (ΔTTI/s) and the associated increase in energy turnover. The rapid drop in NADH autofluorescence is indicative of net NADH oxidation. Similarly, the simultaneously occurring rapid rise in FAD autofluorescence reflects net FADH2 oxidation. The initial minima (and maxima) in the NADH and FAD responses were reached 7.4 ± 0.4 s after the change in stimulation frequency. This value was not different between NADH and FAD or an increase or decrease in stimulation frequency.

Figure 4. Representative example of an experimental run at 3 Hz.

Figure 4

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A, tension. B, tension time integral per second (TTI/s). C and D, NADH and FAD autofluorescence. The arrows correspond to changes in TTI/s (ΔTTI/s), NADH and FAD amplitude during the up transient (from low to high stimulation frequency), or during the down transient (from high to low stimulation frequency).

Early rapid changes in NADH autofluorescence

The amplitudes of the early changes in NADH autofluorescence were linearly related to the changes in TTI/s (ΔTTI/s), both upon an increase in stimulation frequency (the up transient, Fig. 5A) and a decrease in stimulation frequency (the down transient, Fig. 5B). The pooled data showed an excellent correlation between NADH amplitude and ΔTTI/s during the up transient (slope: 0.69 ± 0.09% mm2 mN−1, R2 = 0.833, P < 0.001, Fig. 5A) and down transient (1.20 ± 0.16% mm2 mN−1, R2 = 0.783, P < 0.001, Fig. 5B), albeit with highly significant differences (P < 0.001) in steepness. The intercepts were significantly larger than 0 (1.96 ± 0.59% during the up transient and 1.72 ± 0.92% during the down transient).

Figure 5. Up–down asymmetry in the amplitude of the early changes in NADH and FAD autofluorescence.

Figure 5

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A and B, NADH amplitude plotted as a function of the change in tension time integral per second (ΔTTI/s) for the up transient at 1, 2 and 3 Hz (R2 = 0.833, P < 0.001) and for the down transient (R2 = 0.783, P < 0.001). C and D, corresponding FAD data for the up transient (R2 = 0.660, P < 0.001) and down transient (R2 = 0.689, P < 0.001). The slopes of the regression lines of the up transients (A and C) were significantly smaller than those of the corresponding down transients (C and D, respectively).

The maximal velocity of net NADH utilization was calculated from the maximal velocity attained during the early fast changes in autofluorescence during the up transient and showed excellent correlation with ΔTTI/s (R2 = 0.857, P < 0.001). The maximal velocity of net NADH utilization during the up transient at 3 Hz (78 ± 13 μm s−1) was lower than the maximal velocity of net NADH production during the down transient at 3 Hz (103 ± 19 μm s−1, P = 0.035).

Early rapid changes in FAD autofluorescence

The pooled data showed a significant correlation between FAD amplitude and ΔTTI/s during the up transient (slope: 1.34 ± 0.24% mm2 mN−1, R2 = 0.660; P < 0.001) and during the down transient (1.84 ± 0.31% mm2 mN−1, R2 = 0.689; P < 0.001). The intercepts were significantly larger than 0 (2.10 ± 1.50% during the up transient and 3.51 ± 1.76% during the down transient). Similar to the NADH observations, the maximal velocity of net FAD production related well to ΔTTI/s (R2 = 0.774; P < 0.001). The maximal velocity of net FAD production during the up transient at 3 Hz was lower (16 ± 4 μm s−1) than the net FAD utilization during the down transient (32 ± 5 μm s−1, P = 0.036).

Steady-state NADH and FAD autofluorescence

The time course of the slow changes in NADH and FAD kinetics upon an increase in stimulation frequency, where NADH and FADH2 reduction favours oxidation, consisted of at least two phases and showed mirror-like responses (Fig. 4). The transition between these phases often was rather abrupt and occurred at different time points during the NADH and FAD transients. The final values, averaged over a period of 10 s at the end of 3 min of stimulation at 3 Hz for NADH (98.0 ± 2.3%) and FAD (109.1 ± 4.8%) did not differ significantly from the values before the start of stimulation (at 0.5 Hz).

Mitochondrial respiration

A typical recording at 37°C is shown in Fig. 6A and the mean respiration values for each experimental condition is shown in Fig. 6B. Maximal complex I-coupled respiration (with glutamate, malate, pyruvate and ADP) was 65 ± 6 pmol O2 s−1 mg−1. Maximal complex I- and II-coupled respiration was 227 ± 13 pmol O2 s−1 mg−1. Maximal complex II-coupled respiration (with succinate and rotenone) was 184 ± 12 pmol O2 s−1 mg−1. None of the samples showed a >10% increase in respiratory rate after addition of cytochrome c, indicative for intact outer membrane function. A significant correlation (R = 0.85) was observed between the ADP-stimulated increase in respiration with complex I substrates and the decrease in respiration after blocking complex I with rotenone.

Figure 6. Mitochondrial respiration using complex I and II substrates.

Figure 6

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A, typical example of an experimental recording. First, the complex I substrates G, M and P are injected into the measuring chamber and respiration is supported by proton leak (for final concentrations, see methods). Subsequently ADP is added, supporting complex I-coupled respiration. CC is added to test outer membrane integrity. The addition of S provides a combined complex I- and II-coupled respiration, also called maximal oxidative phosphorylation. ROT blocks complex I function and the respiration observed are supported by complex II only. AA is added to allow a correction for background oxygen consumption. B, mean values for oxygen consumption (JO2 mg−1 s−1) in each of the conditions indicated in (A). AA, antimycin A; CC, cytochrome c; G, glutamate; M, malate; P, pyruvate; ROT, rotenone; S, succinate.

The temperature coefficients (Q10) of complex I- and II-coupled respiration and the total (complex I + II) were, respectively, 2.53 ± 0.29, 1.73 ± 0.10 and 2.07 ± 0.11. The Q10 of complex I- and II-coupled respiration were significantly different (P < 0.05). Because of the fixed stoichiometry between O2, NADH and FADH2, complex I-coupled respiration corresponded to a maximal velocity of NADH utilization of 143 ± 14 μm s−1 and complex II-coupled respiration corresponded to a maximal velocity of FADH2 utilization of 291 ± 19 μm s−1 at 27°C.

Discussion

In this study the kinetics of NADH and FAD autofluorescence were measured simultaneously for the first time together with tension production, on a second-to-second basis in intact cardiac trabeculae. The main findings are: (1) the NADH and FAD transients upon a change in stimulation frequency changed in opposite direction, with rather similar time courses; (2) net velocities of the NADH and FAD supply and utilization were linearly related to changes in the TTI/s and the net NADH/FAD velocity ratios varied in agreement with the predicted 5:1 proportion; (3) the amplitude of early rapid change in the NADH and FAD autofluorescence upon an increase in stimulation frequency was smaller than the amplitude of the change upon a decrease in stimulation frequency (up–down asymmetry); and (4) the maximal velocity of FADH2 utilization in intact trabeculae was considerably smaller than the maximal velocity observed, which is suggestive of a considerable complex II reserve capacity in vivo. These results show that simultaneous measurement of contractile properties and NADH and FAD kinetics in cardiac trabeculae provides additional information on mitochondrial function.

Similarities between NADH and FAD responses

NADH and FADH2 utilization in the electron transport system depend on complex I and complex II activity, respectively. In steady state, both NADH and FADH2 production should by definition match utilization, but the initial changes in NADH and FAD concentration and the time course to a new steady state after an abrupt change in stimulation frequency are not necessarily the same.

Overall, the NADH and FAD responses upon a change in stimulation frequency changed in opposite directions. This is in accordance with the changes observed during ischaemia and reperfusion in intact hearts (Aldakkak et al. 2011). In agreement with previous studies (Brandes & Bers, 1996, 2002), we observed that rapid changes in NADH concentration occurred within seconds after a change in stimulation frequency. The FAD autofluorescence signals showed a similar time course. The initial maxima and minima in NADH and FAD autofluorescence were reached in 7.4 ± 0.4 s after the change in stimulation frequency. No significant differences were observed in the time to the initial maxima and minima of the NADH and FAD transients. These rapid changes are compatible with the rapid increase in oxygen consumption (within 10 s) in intact trabeculae at high stimulation frequency (Van der Laarse et al. 2005).

The early rapid changes in NADH and FAD autofluorescence showed a linear relationship with a change in contractile output (ΔTTI/s). In addition, the maximal velocities of net NADH and FAD utilization and production also correlated well with ΔTTI/s. The maximal velocities derived from the minima and maxima in the differentiated NADH and FAD signal reached during the up transient from 0.5 to 3 Hz amounted to 78 ± 13 and 16 ± 4 μm s−1. These values reflect the net decreases in NADH and FADH2 concentration when their rate of utilization is abruptly increased. It can be noted that our estimates of the velocity of net NADH consumption are compatible with the velocity of NADH production of 0.3 mm s−1 used in the models of Cortassa et al. (2003, 2006) and Hatano et al. (2011).

In our experiments, glucose was used as the substrate. The overall NADH and FADH2 production in the glycolysis and Krebs cycle takes place in a 5:1 proportion. The maximal net velocities of NADH and FADH2 production (and utilization) observed at different stimulation frequencies varied in proportion with the increase in contractile output and are in agreement with this theoretical 5:1 prediction. It can be concluded therefore that our observations indicate that the rapid changes in NADH and FAD concentration are tightly coupled to the changes in contractile output.

Up–down asymmetry in NADH and FAD

The amplitude of the rapid decrease in NADH autofluorescence upon an increase in stimulation frequency (the up transient) was smaller than the rapid increase upon a decrease in stimulation frequency (the down transient). The same holds in the opposite direction for the rapid changes in FAD kinetics. These findings are reminiscent of the on–off asymmetry observed in skeletal muscle (Wüst et al. 2013). It can be noted that the net increases in NADH (and FADH2) concentration during the down transient result from excess NADH (and FADH2) production in glycolysis and the Krebs cycle. The changes in NADH and FAD concentration thus provide a lower estimate of the steady-state rates of NADH and FADH2 production reached just before the decrease in stimulation frequency. Conversely, the changes in NADH and FAD concentration measured upon an increase in stimulation frequency provide a lower estimate of the steady-state rates of NADH and FADH2 utilization reached just before the increase in stimulation frequency. The larger amplitude of the early rapid parts of the NADH and FAD transients during the down transient relative to the up transient indicates a mismatch in the adjustment in the supply and demand of NADH and FADH2. As is illustrated in Fig. 7, the up–down asymmetry in the NADH and FAD responses can be explained by a faster adaptation of demand, a slower adaptation of supply, or both. The physiological implication could be that the early feedback between supply and demand upon an increase in contractile activity is stronger than during a decrease in activity.

Figure 7. Schematic diagram of the velocities of NADH or FADH2 supply and demand upon an increase and decrease in stimulation frequency.

Figure 7

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At t = 0 (s), the stimulation frequency is increased from 0.5 to 3 Hz and at t = 180 (s) the stimulation frequency is reduced from 3 to 0.5 Hz. At t = 0 (s), the velocities of NADH or FADH2 supply and demand (Vs and Vd, respectively) are equal and adapted to the relatively low rate of ATP utilization. Upon the increase in stimulation frequency, the velocities of NADH or FADH2 supply increase to a new steady state. To explain the decrease in NADH and FADH2 concentration upon the increase in stimulation frequency at t = 0 (s), the initial velocity of demand needs to be larger than the initial velocity of supply. At time = tmin or tmax (∼7.4 s), the minima (or maxima) in NADH and FADH2 concentration are reached where Vs equals Vd. The red area indicates the amplitude of reduction in the NADH or FADH2 concentration. The up–down asymmetry in the NADH and FAD transients, i.e. an increase in the blue area, can be explained by either an initially faster adaptation of demand (red dotted line) and/or a slower adaptation of supply (blue dotted line).

It is well established that the in vivo ADP concentration does not vary significantly with changes in workload (From et al. 1986; Balaban, 2006). Moreover, if a rapid imbalance in ADP concentration would be present, its time course would be expected to be the same for an increase or decrease in the rate of ATP utilization. It is therefore very unlikely that differences in ADP concentration could explain the up–down asymmetry. However, asymmetry in intracellular Ca2+ handling possibly associated with a redistribution of Ca2+ stored in the sarcoplasmic reticulum and mitochondria on a relatively fast beat-to-beat basis could serve as an explanation for the up–down asymmetry. Krebs cycle enzyme activity is expected to be higher during the activated state, which is because of Ca2+ activation of glycolysis as well as because of Ca2+ dependence of enzyme activity within the Krebs cycle (Denton, 2009). Similarly, mitochondrial complexes (not only complex I and II, but also others) are activated upon an increase in stimulation frequency, leading to higher velocity of NADH utilization at high stimulation frequency (Gandra et al. 2012; Glancy & Balaban, 2012). Previous results of Bers and colleagues in pyruvate containing solutions (Brandes & Bers, 2002) showed good agreement between the time constants of the slow recovery in NADH fluorescence and mitochondrial Ca2+ concentration and are compatible with this notion. Our results are thus consistent with an important role for the free mitochondrial Ca2+ concentration in parallel activation of the NADH and FAD producing and utilizing pathways, when energy requirements are altered (Chance, 1972; Balaban, 2006; Phillips et al. 2012). In pathological situations where mitochondrial Ca2+ loading is reduced such as in heart failure (Maack et al. 2006), it is possible that activation of Krebs cycle and mitochondrial enzymes is reduced. This could have major consequences for the balance between the NADH and FAD producing and utilizing pathways upon rapid changes in cardiac workload.

Reserve capacity of complex II

Our values for the velocities of net NADH and FADH2 consumption (derived from the differentiated NADH and FAD signals) may be compared with estimates derived from the oxygen consumption using high-resolution experiments in vitro, which represent the maximal activities of complex I and II. The down transient velocities at 3 Hz stimulation (103 ± 19 and 32 ± 5 μm s−1) were larger than the up transient velocities and therefore probably can be considered as our best estimates of the maximal velocities that the NADH and FADH2 utilization reached during steady state. The maximal complex I-coupled respiration measured in the respirometer amounted to 143 ± 14 μm s−1. This value is 72% of the maximal velocity of NADH utilization of complex I. In contrast, the maximal complex II-coupled respiration in vitro corresponded to 291 ± 19 μm s−1, while the velocity of FADH2 utilization in intact trabeculae was only 32 ± 5 μm s−1. This is only 11% of the maximal velocity in vitro. This suggests that complex II activity has a very high reserve capacity in intact trabeculae. The reason for this large difference in reserve capacities of complex I and II is currently unclear.

It can be noted that the temperature coefficient (Q10) of complex I-coupled respiration was significantly higher than that of complex II-coupled respiration. As steady-state NADH and FADH2 supply and demand are matched, this would imply that the difference in complex I and complex II reserve capacity at body temperature is even larger than the difference observed at 27°C.

The maximal oxygen consumption derived from the respirometry measurements in permeabilized right ventricular tissue amounted to 0.23 mm s−1 (at 37°C). The maximal rate of oxygen consumption estimated in electrically stimulated intact trabeculae at 37°C was 0.58 mm s−1 (Van der Laarse et al. 2005) and is almost exclusively used by mitochondria (Gibbs & Loiselle, 2001). This estimate is approximately two times larger than the value obtained in permeabilized tissue. In both cases tissue volume was derived from measurements of the wet weight of the samples. It should be noted that osmotic myofilament lattice expansion of the permeabilized tissue and adhering solution causes wet weight to be at least 30% larger than intact tissue weight. In addition, differences in experimental conditions also complicate this comparison.

Irrespective of the cause of this difference it is clear that considerable reserve capacity exists in complex II activity. Consequently, it is probable that contractile performance will be affected under pathophysiological conditions when mitochondrial complex I is dysfunctional, such as in sepsis or heart failure (Brealey et al. 2002; Wüst et al. 2012). Moreover, this reserve capacity could be functionally relevant when mitochondrial supercomplexes are reorganized to meet altered substrate provision (Lapuente-Brun et al. 2013).

Acknowledgments

Tahe authors would like to thank Henri Tellegen from OceanOptics and Milan Verwoert and Heder de Vries for help during the experiments. Dr Willem van der Laarse is acknowledged for insightful discussions.

Glossary

FAD

flavin adenine dinucleotide

MRC

maximal rate of contraction

MRR

maximal rate of relaxation

NADH

nicotinamide adenine dinucleotide

PMT

photomultiplier tube

Q10

temperature coefficient

Tpeak

maximal tension

TTI/s

tension time integral per s

Additional Information

Competing interests

R.C.I.W. and G.J.M.S. do not have competing interests to declare, M.H. is director of IonOptix LLC (Milton, MA, USA).

Author contributions

All experiments were performed at the VU University Medical Centre (Amsterdam, The Netherlands). R.C.I.W. and G.J.M.S. collected, analysed and interpreted the data; M.H. was involved in the design of the measuring device. All authors approved the final version for publication.

Funding

This work was supported by Netherlands Cardiovascular Research Initiative (CVON2011-11 ARENA).

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