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Address correspondence to Tullio Pozzan, Dept. of Biomedical Sciences, University of Padua, Viale G. Colombo 3, I-35121 Padua, Italy. Tel.: 39-049-827-6070. Fax: 39-049-827-6049. email: tullio.pozzan{at}unipd.it
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Abstract |
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Key Words: calcium; cameleon; in vivo; Na+/Ca2+ exchange; two-photon microscopy
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Introduction |
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As to skeletal muscle, using targeted aequorins, mitochondria in myotubes were shown to accumulate Ca2+ in response to different stimuli (Brini et al., 1997; Challet et al., 2001), and myotubes from dystrophic mdx mice showed an elevated uptake of Ca2+ into mitochondria during potassium-induced cell depolarization (Robert et al., 2001b). There is also indirect evidence that [Ca2+]m increases may serve to adjust ATP production to the physiological demands of muscle and other tissues (McCormack and Denton, 1990; Robb-Gaspers et al., 1998; Jouaville et al., 1999). Furthermore, with using rhod-2, rises of [Ca2+]m after but not during tetanic stimulation were observed in mature isolated skeletal muscle fibers (Bruton et al., 2003a,c); but see Lannergren et al. (2001) for contrasting results. All these studies, valuable as they are, present well-known shortcomings. First, rhod-2 (used generally as an AM-ester) appears to distribute to mitochondria only, or mainly, when applied at low temperature (Trollinger et al., 1997); this limits the use of rhod-2 to in vitro systems in which the incubation temperature can be easily influenced. Second, rhod-2 measures [Ca2+] solely by changes in fluorescence intensity and is therefore prone to the general limitations inherent to nonratiometric dyes, including major artifacts due to movement; this is obviously critical for observations during muscle contraction. Third, the amount of photons emitted by recombinant aequorin is quite low and thus makes the analysis of single cells difficult and with relatively low time resolution. Fourth, all measurements of [Ca2+]m in skeletal muscle during contraction have been performed in in vitro systems using myocytes (Trollinger et al., 1997) or myotubes (Brini et al., 1997; Challet et al., 2001; Robert et al., 2001b); i.e., muscle fibers at a largely immature stage where the kinetics of the Ca2+ changes are far slower than those occurring in vivo (Koopman et al., 2003).
Genetically encoded fluorescent ratiometric Ca2+ sensors (called "yellow cameleon" [YC]) were introduced in 1997 (Miyawaki et al., 1997). Upon binding of Ca2+, they respond with an increase in fluorescence resonance energy transfer (FRET; Miyawaki et al., 1997) and are therefore optimal for experiments with monochromatic excitation and ratiometric read-out, as used in two-photon confocal microscopy. Genetic engineering enables YCs to be specifically targeted to different subcellular locations (such as the nucleus [Miyawaki et al., 1997], ER ER [Miyawaki et al., 1997], and mitochondria [Arnaudeau et al., 2001]) and thus permits the measurement of [Ca2+] therein. Recently, cytoplasmic YCs have been used in living invertebrates (Kerr et al., 2000; Diegelmann et al., 2002; Fiala et al., 2002; Reiff et al., 2002; Ikeda et al., 2003; Shyn et al., 2003; Suzuki et al., 2003), and the zebrafish (Higashijima et al., 2003); surprisingly, YCs have yet to be shown to function correctly in live mammals.
We transfected mouse hind-limb muscle in situ with a second generation YC (YC2; Miyawaki et al., 1997) targeted to the cytoplasm or mitochondria, and monitored [Ca2+]c and [Ca2+]m by two-photon microscopy. The initial part of our study sought to validate YC2 as a reliable Ca2+ indicator and the data obtained show that cytoplasmic YC2 is capable of recording short-lived rises in [Ca2+]c in the millisecond range, and correcting for gross movement artifacts. Our subsequent studies to address [Ca2+]m homeostasis demonstrated that the rise in [Ca2+]m observed during single twitches and tetanic stimulation in live mammalian skeletal muscle is synchronized with the physiological induction of muscle contraction.
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Results |
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Discussion |
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For the transient expression of YC2 in mouse tibialis anterior muscle, we adopted an established electroporation-based protocol (Dona et al., 2003), with minor modifications. This technique permits the observation of YC2 fluorescence up to 3 wk after transfection (Fig. 1, A and B); i.e., long after cessation of any inflammatory responses and muscular repair mechanisms due to the operation. Indeed, the transfected muscles exhibited no obvious signs of damage or inflammation when prepared for microscopic visualization. This type of gene transfer provides an elegant and powerful way to study muscle physiology with genetically encoded probes, circumventing typical problems of other genetic manipulation techniques. First, it is fast, easy to perform, and shows a high transfection efficiency. Second, it is absolutely specific for the electroporated region and therefore is not expected to cause systemic interferences. Third, because it is applied only few days before microscopic examination, also ontogenetic disturbances are absent.
Using stimulation frequencies ranging from 1 to 50 Hz, we show that the mean [Ca2+]c in mammalian skeletal muscle in situ (as measured both by YC2 and split YC2) is in direct correlation to the intensity of stimulation (Fig. 3, A and B). Regarding the correlation shown in Fig. 3 B between stimulation frequency and YFP/CFP ratio, this is in agreement with work previously published for isolated mouse muscle fibers loaded with indo-1 (Helander et al., 2002). It is worth stressing that these results primarily reflect the number of twitches captured at the different stimulation frequencies (Fig. 3 A), although they do not provide any information about the ratio amplitude of each individual twitch. These measurements are of interest, nonetheless, because they provide an estimate of the mean cytosolic Ca2+ level in a period of several seconds, a potentially important parameter regarding the regulation of other Ca2+-dependent processes, such as the control of gene expression (Dolmetsch et al., 1998; Li et al., 1998). Evidence for the robustness of the method used in our studies is the fact that a Ca2+-insensitive construct similar to YC2 did not report any changes in ratio signal (Fig. 3 B). This clearly indicates the reliability of the YC2 read-out under conditions of strong movement of the observed object.
In isolated mouse extensor digitorum longus (EDL) muscle and soleus fibers, using furaptra, the onset of [Ca2+]c rise and the beginning of contraction are almost concomitant, whereas the maximum of contraction takes place typically several tens of milliseconds after the Ca2+ peak (Baylor and Hollingworth, 2003). The latter is reached at nearly physiological temperatures in 3 to 5 ms. Taking advantage of the scanning process of the confocal microscope, we could obtain good time resolution of the kinetics of both contraction and [Ca2+]c rise, as monitored by YC2. As demonstrated by Figs. 4 B and 6, the peak of the YC2 ratio change is reached with a significant delay with respect to that revealed by a fast Ca2+ dye such as furaptra (Baylor and Hollingworth, 2003); indeed, the peak of [Ca2+]c as revealed by YC2 and maximum contraction seem to coincide. We have not attempted to analyze in detail the reasons for this relatively slow response of YC2 (a clear limitation for the detailed kinetic analysis of cytoplasmic Ca2+ handling in muscle tissue), because this study does not delve into the kinetics of [Ca2+]c versus that of contraction; rather, our main concern was to establish that relatively fast [Ca2+] transients (within 10 to 50 ms) are capable of being detected by YC2, even in conditions of substantial specimen movement. Regarding the decay phase, at low frequency stimulation (Fig. 4 B, 110 Hz), the YFP/CFP ratio returned to baseline values within milliseconds, practically in synchrony with contraction. In contrast, at 50 Hz, the recovery to basal [Ca2+]c lasted some seconds (Fig. 2). As judged from the concise response of YC2 upon single twitches (Fig. 4 A, see 1 Hz and 5 Hz ratio images), this long-lasting recovery seems to reflect a true process, though at present some artifact due to an intrisic delay of the probe cannot be completely excluded (for example see Kerr et al., 2000). Regarding the peak maxima shown in Fig. 4 B, ratio values reach 1.27 ± 0.02 (mean ± SEM, n = 10) at the point of maximum contraction when using stimulation frequencies of 110 Hz. At 50 Hz, a slightly higher peak value of 1.32 ± 0.02 (mean ± SEM, n = 10) was observed, correlating with what is known about muscle physiology.
As to mitochondrial Ca2+ accumulation, to our knowledge, there is no data until now concerning what happens in the organelles of mature muscle fibers during contraction itself; furthermore, data obtained seconds after suspension of stimulus are contradictory (Lannergren et al., 2001; Bruton et al., 2003c). It should also be noted that the in vitro studies on mature fibers presented so far are based on rhod-2 fluorescence measurements, well-known to be prone to movement artifacts (Lannergren et al., 2001; Bruton et al., 2003b,c). At present, the only data available concerning the behavior of [Ca2+]m during skeletal muscle contraction are those obtained in myocytes (Trollinger et al., 1997, 2000) and myotubes (Brini et al., 1997; Challet et al., 2001; Robert et al., 2001b), but their relevance with respect to the behavior of the mature fibers is debatable. Using mitochondrially targeted YC2, we show here for the first time that mitochondria in mouse skeletal muscle observed in situ and under highly physiological conditions do indeed take up Ca2+ both during tetanic contraction (Fig. 5, K and L) and single twitches (Fig. 6 B). In terms of amplitude, assuming that the affinity of mitochondrially located YC2 is the same as that observed in the cytosol, the peaks in [Ca2+] are somewhat smaller in mitochondria (mean 1.14 ± 0.02 SEM, n = 21 videos, 150 contractions, six experiments) compared with the cytosol (mean 1.26 ± 0.03 SEM, n = 18 videos, 97 contractions, three experiments). During tetanic stimulation no obvious major differences were noticed in the kinetics of rise and decay of the mitochondrial Ca2+ signal compared with those in the cytosol, although when the phenomenon was analyzed at higher temporal resolution (as in the single twitch experiments) mitochondria generally showed some delay, in the order of 10 ms, both in the beginning of the Ca2+ rise and in reaching the peak when compared with the cytosol (Fig. 6). Similarly, the decay phase is slightly slower in mitochondria. We would like to stress that whereas the fast Ca2+ uptake rate could have been predicted (particularly now that the kinetic properties of the mitochondrial uptake system have been unraveled by direct electrophysiological measurements by Kirichok et al., 2004), the speed of the decay phase, completed in a period of 100 ms, is much faster than it could have been anticipated from in vitro measurements. To get some insight into how this fast expulsion of Ca2+ may occur, we tested its kinetics in the presence of an inhibitor of the Na+/Ca2+ exchanger. Our data show that under these conditions the decay in [Ca2+]m is slower than in the control (Fig. 6, B and C), suggesting an important role of the Na+/Ca2+ exchanger for this process.
A final important observation is obtained by the experimental protocol of Fig. 5 L. Here, we used a stimulation protocol that is similar to that physiologically delivered by fast motor neurons to their target fibers, i.e., bursts of high frequency (50 Hz) and short temporal extension (500 ms; Hennig and Lomo, 1985). From the data presented in Fig. 5 L (upper panel), it seems that under these conditions [Ca2+]m remains elevated within the mitochondrial matrix far longer than the stimulus itself and if another train of stimuli is delivered at short distances, i.e., within 2 to 3 s, [Ca2+]m remains continuously elevated above basal level. In contrast, in the cytoplasm under the same experimental regime the basal value is achieved almost instantaneously after cessation of the stimulus (Fig. 5 L, lower panel). Although the degree of elevation varied between the different fibers, and we cannot exclude an inherent slow of rate of the YC2 probe, our data suggest that under conditions similar to a strong physiological stimulation of the muscle, the full power of the Ca2+-dependent mitochondrial dehydrogenases should be available, thus permitting the most efficient use of oxidative substrates and a maximal speed of ATP synthesis.
In summary, the data obtained with mammalian skeletal muscle indicate that (a) in vivo transfection using electroporation is a powerful method for expressing second messengersensitive probes in the muscle of a living mammal, (b) cytoplasmic YC2, in spite of some kinetic limitations, is a useful reporter of changes in [Ca2+]c during contraction and relaxation, and (c) the mitochondrially targeted YC2 variant reveals a rise in [Ca2+]m during contraction, not only during tetanic stimulation, but also in single twitches.
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Materials and methods |
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Animals
For most of the shown experiments, C57BL/10 mice with an age of 612 mo were used; for calibration experiments, SV159 mice were also used; all animals were obtained from Charles River Italia. Use and care of animals was approved by the local authority for veterinary services and followed the guidelines of the Italian Health Ministry (Law 116/92 and Art. nr. 8/94). An i.p. injection containing a combination of Rompun® (Bayer) and Zoletil 100® (Laboratoires Virbac) was used for anesthesia.
Transfection method
For the heterologous expression of YCs, an electroporation-based transfection method of the tibialis anterior muscle was used very similar to the one described recently (Dona et al., 2003). In brief, the shaved skin of the left lower hind limb of the anesthetized animal was opened at the border between anterior and posterior hind limb muscles with a longitudinal cut of 1 cm length. The fascia was cut and an electrode was inserted between muscle and bone. With a modified 100-µl microsyringe (Exmire Microsyringe; ITO Corp.), 30 µg of expression plasmid were injected into the exposed tibialis anterior muscle before placing the second electrode on the muscle. With an Electro Square Porator (model ECM 830; BTX Genetronics), five pulses of 20 ms duration were applied with a voltage of 25 V. The electrodes were removed, and the wound was closed and sterilized. The antibiotic Terramicina® (Pfizer Italia) was injected as antiinflammatory prevention.
In vivo two-photon microscopy
Preparation of muscle and nerve.
23 wk after transfection the operated animal was anesthetized and the area around the original incision shaved. The sciatic nerve was isolated and a custom-made silver-microelectrode was inserted for subsequent stimulation of muscle contraction. The distal tendon of the transfected tibialis anterior muscle was severed and the fascia removed. The mouse was then transferred onto a custom-made plastic table, the muscle was fixed with a surgical clamp attached to the severed tendons, and the complete assembly mounted for observation.
Microscopy.
A multi-photon system (Radiance 2100 MP; Bio-Rad Laboratories), equipped with a Tsunami mode-locked, tunable, femto-secondpulsed Ti/sapphire laser, optically pumped by a Millennia VsS 5W green laser (both from Spectraphysics Lasers), was used for two-photon microscopy. The laser output is capable of generating 100-femto-second pulse trains at a rate of 82 MHz. The excitation wavelength was 830 nm, controlled with a spectrometer (Ocean Optics USB2000; getSpec). The microscope (Eclipse E600FN; Nikon) was equipped with a Leica objective (PL APO 63x/1.2W CORR; Leica); an optical gel (0.2% polyacryl; Viscotirs Gel®; Ciba Vision) was used as immersion medium. A Direct Detection System (Bio-Rad Laboratories), fitted with a 500LP DC dichroic mirror and HQ450/80 and HQ535/50 emission filters (Chroma Technology Corp.), was used for the simultaneous detection of CFP and YFP emission signals, respectively. Typically, gain and offset of the photomultipliers were kept constant while laser intensity was adjusted according to the depth of the observed fiber; only fibers from the first to the second outer layer (i.e., up to 100 µm in depth) were imaged. The LaserSharp2000 software package (Bio-Rad) was used for data acquisition. Unless otherwise indicated, video microscopy was performed at 128 x 128 pixels per image, with a 3.95-Hz acquisition frequency. For colocalization studies, images were acquired at 1,024 x 1,024 pixels, using a 560DCLPXR dichroic mirror; emission filters E560SP and HQ600/50 were used for the detection of YC2 and TMRM, respectively. Calcium levels were monitored in the cytoplasm with YC2 or split YC2, and in mitochondria with 2mtYC2. Co-expressed CFP-CaM and YFP in the cytoplasm, or mtCFP and mtM13-YFP in mitochondria, served as controls, and did not show an increase in YFP/CFP fluorescence intensity ratio during muscle contraction; in the controls for mitochondria, a decrease in ratio was sometimes observed during contraction. The latter effect is most likely due to the following: in kinetic experiments, the high image acquisition rate imposes a limited spatial resolution; individual mitochondria are thus not resolved and background correction becomes defective; because the background level is higher in CFP images, there is an artificial enhancement of the CFP signal during contraction.
Induction of muscle contraction.
For the stimulation of muscle contraction, a custom-made electrode was attached to the sciatic nerve and connected to an adjustable stimulator (Scientific Research Instruments). The typical voltage applied was in the 0.10.2-V range, with variable stimulation frequencies, as indicated in the text; each pulse had a duration of 5 ms. The stimulation intensity was adapted to the lateral movement of the fiber during contraction in the microscopic field.
Injection of probes.
500 nM TMRM, 10 µM FCCP, and 200 µM CGP37157 were injected locally into the observed muscle with a 30-gauge needle (Artsana) with a typical volume of 5060 µl. To allow the muscle to recover from the injection-induced swelling, microscopic observation was interrupted for 25 min.
Cryopreservation and sectioning of muscle
After imaging, the complete anterior tibialis muscle was removed, washed in physiological solution, incubated for 10 min in relaxation buffer containing (in mM): 100 KCl, 5 EGTA, 5 MgCl2, 3 2,3-butanedione monoxime, 0.25 dithiothreitol, 10 histidine, pH 7.8, and then fixed for 2 h at 4°C in PFA solution (4% wt/vol paraformaldehyde, 4% wt/vol sucrose in PBS, pH 7.4). After washing in distilled water, the fixed muscles were then snap-frozen in isopentane cooled in liquid nitrogen (Wu et al., 1985), and stored at 80°C. For sectioning, muscles were embedded according to standard procedures in Jung Tissue Freezing Medium (Leica) at 20°C. A microtome (model CM1850; Leica) was used to prepare 10- and 30-µm-thick sections (for transversal and longitudinal sections, respectively), which were placed on electrostatically charged microscope slides (Superfrost Plus; BDH Laboratory Supplies). Transmission and fluorescence images were acquired using either a microscope (model BX60 equipped with a 50W HBO mercury arc lamp, a PLAN2x/0.05 objective, and a DP50 camera; Olympus), using Photoshop 6.0 (Adobe Systems), or a Bio-Rad Laboratories confocal microscope (Radiance 2100 MP equipped with an argon laser for 488 nm excitation of cameleon fluorescence, a Nikon 60x/1.4 Plan Apo objective, a 500 DCLPXR beamsplitter, and a HQ515/30 emission filter; all filters were from Chroma Technology Corp.); confocal images were taken at 1,024 x 1,024 pixel resolution with LaserSharp 2000 software (BioRad).
Data analysis and ratiometric videos
Analysis of the two-photon in vivo imaging data was performed using the public domain ImageJ program (developed at the United States National Institutes of Health by Wayne Rasband and available freely on the Internet at http://rsb.info.nih.gov/ij/). For the calculation of YFP/CFP fluorescence intensity ratios, the following procedure was used: the fluorescent fibers were outlined as regions of interest (ROI) by thresholding the YFP image sets; the mean greyscale values of the ROIs were calculated in both YFP and CFP image sets; the obtained values were transferred to Excel 2002 (Microsoft), where all subsequent calculations and graphs were performed. Ratio plots were normalized using as reference the average value of the images before the first stimulation (because laser illumination causes an initial photo-isomerization of YFP; Miyawaki et al., 1997; Filippin et al., 2003), the average used for normalization considered only images after a plateau of the YFP/CFP ratio was reached). The kinetics of the YFP/CFP ratio during twitches in the cytosol (as shown in Fig. 4 B) was determined by measuring ratio values along six parallel lines within the fiber, thus generating average ratio intensities along the fiber; the average ratio intensities along the fiber were "aligned" to the beginning of the first twitch observed in the respective image. For each stimulation frequency, the same procedure was performed on different images (n = 10). The kinetics of the YFP/CFP ratio during twitches in the cytosol and mitochondria (as shown in Fig. 6) were determined by the fluorescence intensities of rectangular ROIs on the respective ratiometric videos: these ROIs had a width of five lines (corresponding to 10 ms in real time), the height of the fiber diameter, and scanned the entire fiber length; alignment was performed as described above. Ratiometric videos were generated in ImageJ using the following scheme: the original 8-bit videos were median filtered (1-pixel kernel), and a pixel-by-pixel ratio image generated using a custom-made plugin (RatioPlus; available freely at the ImageJ web site). The resulting 32-bit greyscale images were pseudo-colored using a rainbow-like look-up table where low and high values are represented by dark blue and intense red, respectively. For visualization only, the pseudo-color ratio images were intensity calibrated: the brightness of each color (representing a specific ratio value) was adjusted based on the fluorescence intensity of the original YFP and CFP images. Unless otherwise indicated, ratio values refer to YFP fluorescence intensity divided by CFP fluorescence intensity.
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Acknowledgments |
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The work of R. Rudolf has been funded by a Marie-Curie fellowship (HPMF CT200201543.01), a grant from the University of Padua, and from the Deutsche Forschungsgemeinschaft (RU 923/1-1). This work was also partially supported by Italian Telethon (grant 1226), from the Italian Association for Cancer Research, from the Italian Ministry of University (Special Projects Prin RBNE01ERXR-001 and 20033054449-001 and FIRB 2002053318-005), from the Italian Ministry of Health, and from the CARIPARO foundation to T. Pozzan.
Submitted: 18 March 2004
Accepted: 1 July 2004
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