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Investigating Second Messenger Signaling In Vivo
C H A P T E R
N I N E T E E N
Investigating Second Messenger Signaling In Vivo ¨diger Rudolf,* Mathias Hafner,† and Marco Mongillo‡ Ru Contents 1. Introduction 2. Multimodal Imaging in Living Mouse Skeletal Muscle 2.1. Ad hoc probe introduction: Injection, electroporation 2.2. Preparation of the mouse for imaging: Time for add-ons 2.3. In vivo imaging of mouse skeletal muscle: Space limitations 2.4. Data extraction 3. Adaptations to Tissues Other Than Skeletal Muscle 3.1. Isolated heart 3.2. Brain imaging—A brief teaser 4. Concluding Remarks Acknowledgments References
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Abstract All known second messengers are small molecules without complex structural features. However, each of them can mediate different, very specific cellular responses upon arrival of distinct extracellular cues in one and the same cell. From this follows the question of how signal specificity is achieved in space and time. Recent work showed that three factors play major roles in determining signal specificity: intensity, pattern, and compartmentalization of signals. Thus, for understanding the signaling in any specific context, generic information on the involvement of second messenger pathway(s) is insufficient and only the precise, time- and space-resolved monitoring of these processes will lead to meaningful insights. Even more demanding, many second messenger-based signaling events can only occur using tissue-specific morphological arrangements (e.g., striated organization of muscle and synaptic specializations in neurons), making the visualization of intact tissue mandatory. Finally, to * Institut fu¨r Toxikologie und Genetik, Karlsruhe Institute of Technology, Hermann-von-Helmholtz-Platz 1, Eggenstein-Leopoldshafen, Germany Institut fu¨r Medizintechnologie der Universita¨t Heidelberg und Hochschule Mannheim, Mannheim, Germany { Venetian Institute of Molecular Medicine, Via Orus 2, Padua, Italy {
Methods in Enzymology, Volume 505 ISSN 0076-6879, DOI: 10.1016/B978-0-12-388448-0.00027-9
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2012 Elsevier Inc. All rights reserved.
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appreciate the physiological relevance of second messenger signals, multiparametric readouts are generally needed. This chapter illustrates modes of multiparametric analysis of signaling in live mouse skeletal muscle and briefly discusses possible applications of the technique in other tissues including heart and brain.
1. Introduction Second messengers mediate an enormous spectrum of cellular responses to external stimuli, ranging from the regulation of cell proliferation and metabolism to cell death. Notably, different signals might arise within one cell at the same time, rendering it absolutely necessary for the cell to achieve signal specificity (Zaccolo and Pozzan, 2003; Zaccolo et al., 2002). Clearly, a vast proportion of this specificity is due to amplitude, pattern (e.g., oscillations), and subcellular compartmentalization of signals. In particular, the latter is completely cell-type dependent and can usually not be reconstituted in vitro in a reliable manner. Therefore, the physiological roles of second messenger signaling can often only be understood when studied in the living tissue or in the intact organism. Since the discovery of green fluorescent protein (GFP), numerous fluorescent, genetically encoded sensors able to monitor second messengers in living cells were created and new ones are continuously developed (Pozzan et al., 2003; Whitaker, 2010; Zhang et al., 2002). In combination with adequate approaches, genetically modified organisms, and molecular or electronic biosensors measuring additional biophysical parameters, the use of GFP-based sensors can now deliver invaluable insights into many second messenger-dependent physiological processes in intact tissue. In most cases, probes for second messenger dynamics exploit Fo¨rster resonance energy transfer (FRET), usually between cyan and yellow fluorescent proteins (FPs), to transform conformational changes in the sensor molecule into measurable changes in their fluorescence spectra. The investigation of such probes in living samples has been enabled by multiphoton imaging, which allows excitation of short-wavelength dyes deep in tissue. However, the merit of tissue penetration of multiphoton light is dampened by its broad excitation range, which makes the discrimination of more than two fluorescent probes difficult. However, since dyes with longer wavelengths can efficiently be visualized even with standard confocal microscopy deep in tissue, multimodal imaging, using multiphoton together with single-photon confocal microscopy is useful to combine the visualization of second messenger signals with other readouts. Transgenic techniques have rendered possible the introduction of genetically encoded probes into mice and a few other model organisms.
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However, at least in mammals, these methods cannot cope with the speed of novel sensors development and the progress in biomedical research. Therefore, faster approaches to introduce molecular sensors into target tissues are needed. While for some organs viral vectors might be ideal, electroporation-mediated gene transfer has proven to be an efficient means in skeletal muscle. This chapter first describes the steps from genetic manipulation of skeletal muscle to the multimodal imaging and data analysis of second messenger signaling and other factors in this tissue in living mice. Then, we introduce adaptations of this paradigm to other tissues, namely, cardiac muscle and neuronal tissue.
2. Multimodal Imaging in Living Mouse Skeletal Muscle Second messenger monitoring alone can be used to investigate the effect of ligands or trigger stimuli on a selected second messenger pathway. However, to understand the physiological meaning of such a signal, second messenger monitoring needs to be coupled to additional readouts. This can be done by combining in vivo imaging with electronic biosensors, such as force transducers (Allen et al., 2011) or patch clamp devices. Alternatively, if fluorescent long-wavelength markers are available, these can be analyzed by using single-photon confocal microscopy in combination with second messenger analysis of FRET-based probes in the multiphoton mode (Ro¨der et al., 2009, 2010). Additional combinations, for example, with second harmonic generation imaging can be envisaged. The ability of multimodal imaging to interrogate simultaneously intracellular signals and functional outputs makes a clear advantage over the single modal analysis. In the following paragraphs, the steps from probe introduction to multimodal imaging and data analysis are exemplified for mouse hindlimb muscle.
2.1. Ad hoc probe introduction: Injection, electroporation Due to their accessibility, muscles of the mouse distal extremities are particularly amenable to probe injection and electroporation (Figs. 19.1A and B and 19.2A–G). On the one hand, injected probes might be cell-permeant or cell-impermeant, such as tetramethyl rhodamine methyl ester (TMRM) (a mitochondrial membrane potential marker; Romanello et al., 2010) or a-bungarotoxin coupled to chromophores (marking AChRs in neuromuscular junctions; Ro¨der et al., 2008, 2010), respectively. On the other hand, electroporation (Figs. 19.1B and 19.2A–G) is a quick and efficient method to introduce cDNAs for expressing genetically encoded fluorescent probes or
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Figure 19.1 Schematic of in vivo-imaging procedure. Typical experiments involving live imaging of mouse skeletal muscle are divided into at least three phases. First, muscles are prepared by probe and/or cDNA injection (A). Second, in many cases, muscle transfection is used to incorporate sensor-encoding cDNAs and/or muscle function-modifying constructs (B). Third, a couple of days later, in vivo imaging is performed, often using multiple excitation and emission modes to gather multifactorial readout data (C). In some cases, additional treatments (e.g., pharmacological or surgical interventions) are adopted (not shown).
other genes or siRNAs, which modulate any tissue function of interest. As shown previously, transfected cDNAs remain actively transcribed and translated for up to 4 weeks after electroporation in skeletal muscle (Dona et al., 2003). This is most likely due to the postmitotic status of adult skeletal muscle. Although very high levels of up to 80% of transfected fibers might be reached
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Figure 19.2 Setups and procedures for transfection and in vivo imaging of mouse skeletal muscle. (A–G) Setup and procedure of muscle transfection. A setup composed of electroporator, electrodes, heated operation platform, and surgical instruments is used (A). Electroporation involves shaving of the lower hindlimb (B), a longitudinal cut to get access to the hindlimb muscles (C), insertion of a spatula electrode (D), injection of cDNA (E), application of the second electrode and electropulsing (F), and closure of the wound by surgical stitches (G). (H–N) Preparation for in vivo imaging. Ten days after electroporation, the wound should be completely healed (H) and the muscle exhibit a pale rosy color (I). After removing the skin above the electroporated muscle (here: tibialis anterior), the transparent epimysium covering the muscle is eliminated (J). Using forceps and scissors, the distal tendon is cut (K, L) and the mouse transferred to a custom-made table for microscopy (M). Using a hemostatic clamp and plastic and cork supports, the distal tendon is then fixed over plastic tubing (N). (O–Q) Transfer to microscope and in vivo imaging. A confocal microscope equipped with an upright stand, standard and two-photon lasers, and water immersion objectives adapted for use without coverglass is used (O) and the custom support is mounted on the microscope object table (P, Q).
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for some constructs, expression of heterologous proteins being targeted to specific sites or organelles, such as the synaptic membrane, mitochondria, or the sarcoplasmic reticulum can be much more cumbersome. This is particularly true for transmembrane proteins of the synapse, since the secretory pathway mediating the delivery of synaptic proteins is scarcely developed in skeletal muscle. For two main reasons, we usually wait a couple of days (10) before imaging transfected muscles. First, a mild, unavoidable inflammation typically occurring at the site of electroporation is completely healed by then. Second, many proteins in skeletal muscle exhibit low turnover rates, such as the AChR, which has a half-life of >10 days. It is also critical to have always the same, experienced persons performing the electroporation, because this strongly reduces damaging of muscle, which is almost zero in the optimal case, but can be problematic upon inadequate performance. It is worth noting that electroporation efficiency might vary from mouse strain to mouse strain and it is particularly low in muscles with a lot of fat and connective tissue, such as in dystrophic animals. Of the different protocols available for muscle transfection, we prefer to use the method from Dona et al. (2003), where the transfected muscle (e.g., tibialis anterior (TA)) is exposed by simple surgery (Figs. 19.1A, B and 19.2A–G). This is ideal for local and muscle-specific transfection and allows transfecting either the whole muscle (by injecting at different sites) or only superficial fiber layers (sufficient for imaging). For transfecting lower hindlimb muscles, such as TA or extensor digitorum longus (EDL), we use spatula electrodes of 1 mm thickness and 4 mm width (Fig. 19.2D). These are connected to a BTX ECM830 square pulse generator (Fig. 19.2A). After orienting the anesthetized mouse on the side to be transfected, the lower hindlimb is stretched over a pillow and crepe tape is used to fix the foot (Fig. 19.2B). After shaving the leg, a longitudinal cut of 6 mm length opens the midline of the lower hindlimb exposing a view on the whitish border between anterior and posterior muscles (Fig. 19.2C). The shiny fascia (see Fig. 19.2J) enclosing this whole arrangement is then opened by a second longitudinal cut. This allows separating anterior from posterior muscles and seeing the tibia. Now, a closed pair of small surgical scissors penetrates between the tendon of the TA and the tibia. Careful opening of the scissors creates a hole through which the electrode can be inserted (Fig. 19.2D). After fixing the electrode on the pillow with tape, a Hamilton syringe is used to slowly inject up to 20 ml of cDNA in physiological solution (less is better, Fig. 19.2E). Depending on the construct, 0.5–20 mg of cDNA might be used. Make sure that no liquid is leaving the injection site, because this usually results in low transfection efficiency. If necessary, additional, cell-impermeant labels should be injected subsequently. Then, carefully place the second electrode on top of the muscle (Fig. 19.2F) and apply pulses in the following pattern: five pulses with 5 ms duration each are given at an interval of 200 ms between pulses. Voltage is
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typically 50 V/cm of muscle. Make sure to adapt the voltage to muscle thickness. Then, carefully remove both electrodes and close the skin with two to four surgical stitches (Fig. 19.2G), followed by sterilizing the wound. Experienced operators perform a transfection in a few minutes, and animals immediately walk, jump, and behave normally after complete recovery from anesthesia. To help fast recovery, let the animals wake up on a heating plate at 37 C. Animals, which do not behave normally few hours after transfection, suffer from pain or infection and should additionally be treated with an antibiotic and analgesic. 2.1.1. Instrumentation and disposables needed 2.1.1.1. Instruments BTX ECM830 square wave pulse generator Two spatula electrodes and cables Heating plate set to 37 C Hamilton syringe (20 or 100 ml) Straight 9-cm surgical scissors Dumont #7 curved shanks forceps (11.5 cm) with and without serrations Narrow pattern curved forceps (12 cm) Surgical needles (size 2) Needle holder (12.5 cm) Surgical silk (size 5/0) Surgery plate for small animals Custom-made pillow to sustain leg for transfection 2.1.1.2. Solutions
Physiological solution to moisten the muscle during the transfection procedure Anesthetics cDNA and other dye solutions in sterile physiological solution 2.1.1.3. Disposables
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2.2. Preparation of the mouse for imaging: Time for add-ons A few days after transfection, the surgical wound should be completely healed (Fig. 19.2H), and when opening the skin of the anesthetized mouse for imaging, the transfected muscle should exhibit a nicely smooth surface and a pale rosy color (Fig. 19.2I). Any sign of inflammation or hematoma indicates severe muscle damage and is strongly counterindicating the use of
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such a specimen for imaging. Healthy muscles can be further processed for in vivo imaging. Depending on the experiment, muscles can again be injected with fluorescent dyes (Ro¨der et al., 2008, 2010; Romanello et al., 2010; Valkova et al., 2011), or they can be attached to force transducers (Allen et al., 2011), muscle electrodes (Rudolf et al., 2004, 2006; Tothova et al., 2006), or other electronic devices. Anesthesia might be adapted to specific needs, but for imaging, we have made the best experience using intraperitoneal injection of a mixture of Zoletil 100 and Rompun. Compared to other anesthetics, there is much less spontaneous movement of muscles, and anesthesia of the animal is deep and reliable for a time period of about 2 h. Also, orienting and fixing the observed muscle properly is crucial. We detach the distal tendon of the TA (Fig. 19.2K and L) and fix it either with a surgical hemostat or hook it with surgical silk to a force transducer in an angle of 70 –80 from the tibia and horizontal to the surface of the support (Fig. 19.2M and N). This makes the muscle more rigid for imaging and further reduces breathing-induced muscle movements. Before detaching the tendon, carefully remove the fascia directly surrounding the muscle (epimysium, Fig. 19.2J). If electroporation was carried out properly, this layer should still be intact, and after a careful longitudinal cut from the distal tendon to the knee, the two halves of the epimysium can be flipped to both sides of the muscle. Make attention to moisten the muscle frequently with physiological solution. In case of muscle force measurements, the tibia has to be properly fixed to the support using hooks. For pure imaging, the use of crepe tape placed over different points of the leg and the body is sufficient. It is ideal to use a custommade metal support with threads at different positions (Fig. 19.2M) to fix the table after mounting of the animal on the microscope table and to position further tools, such as micromanipulators, force transducers, etc. 2.2.1. Instrumentation and disposables needed 2.2.1.1. Instruments Custom-made support for mounting the animal onto the microscope object table Straight 9-cm surgical scissors Dumont #7 curved shanks forceps (11.5 cm) with and without serrations Narrow pattern curved forceps (12 cm) Hartman straight hemostat (10 cm) for fixing distal tendon to metal support Surgery plate for small animals 2.2.1.2. Solutions
Physiological solution to moisten the muscle during the transfection procedure Anesthetics: Zoletil 100 (Virbac) and Rompun (2% xylazine, Bayer) Dye solutions in sterile physiological solution (if applicable)
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2.2.1.3. Disposables
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2.3. In vivo imaging of mouse skeletal muscle: Space limitations Once mounted on the metal support, the mouse is transferred to the microscope for imaging. Use an upright microscope, which permits working both in single-photon confocal as well as multiphoton mode (Fig. 19.2O). The object table of the microscope should permit application of your custom-made support carrying the prepared mouse and electronic z-plane motion (Fig. 19.2P). At its lowest position, the distance between object table and objective front lens should be at least 5 cm. You will need that space for mounting all the accessories on top of the table. We use a Leica DMRE TCS SP2 microscope equipped with acousto-optical beam splitter (AOBS), three photomultiplier tubes for single-photon detection, two non-descanned detectors for multiphoton detection, and 20/0.7 (Fig. 19.2A) and 63/1.2 water immersion objectives corrected for use without coverglasses (Fig. 19.2O). As a general remark, always take care to use well-corrected objectives with numerical apertures as high as possible. This will enormously impact on your image quality and the amount of light needed for imaging. To directly couple the objective to the tissue of interest, we use artificial tear gel, which has similar optical properties as water, but does not easily drain off due to its high viscosity. Using such setup, the analysis of many different parameters in the living muscle can be executed. Limitations come from three main sources: probe availability, imaging speed, and spectral overlaps. But developments in probe generation and microscope technology continuously reduce the range of these limits. Due to the versatility of the approach, we can here only exemplify how multimodal imaging leads to more meaningful insights. We recently described that the localization of protein kinase A type I (PKA I) in close proximity to a subsynaptic cAMP microdomain is crucial for the proper turnover of acetylcholine receptors at that site (Ro¨der et al., 2010). To address this issue, we needed to correlate cAMP signals with synapse morphology and the presence of PKA I. Therefore, we transfected a FRET-based genetically encoded fluorescent cAMP sensor (RIa-EPAC), which also acts as an in vivo marker for PKA I localization. Ten days after transfection, we additionally injected the synaptic marker, a-bungarotoxin, coupled to the infrared dye, AlexaFluor 647. Then, using single-photon microscopy (Fig. 19.3A–D), we determined PKA I localization and synapse morphology. This showed the presence or absence of PKA RI in different synapses (Fig. 19.3A–C) and the size of these synapses in 3D (Fig. 19.3D). Subsequent multiphoton imaging (Fig. 19.3F–G) permitted to investigate the cAMP signaling in these fibers. Figure 19.3E shows one synapse
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Figure 19.3 Example for multifactorial in vivo imaging. Mouse tibialis anterior muscles were transfected with RIa-EPAC, a FRET-based sensor for cAMP that targets to a PKA RIa-specific microdomain in close proximity to neuromuscular junctions. Ten days after transfection, muscles were injected with fluorescently labeled a-bungarotoxin, a marker for acetylcholine receptors in neuromuscular junctions, and then imaged in vivo using confocal (A–E) and two-photon imaging (F, G). (A) Maximum z-projection of a 135 mm thick z-stack of optical slices taken at 3 mm interslice distance. The scale bar represents 50 mm. YFP-fluorescence signals of RIa-EPAC are shown in A2, a-bungarotoxin staining in A1. Individual optical sections of the boxed regions are shown in B and C. (B, C) Details of the optical section numbers 22 (B) and 28 (C) from the maximum z-projection depicted in A. The scale bar represents 50 mm. (B1 and C1) a-bungarotoxin staining signals. (B2 and C2) RIa-EPAC fluorescence signals. (D) Quantification of volumes of neuromuscular junctions a, b, and c. (E) Maximum z-projection of one synapse in single-photon mode. The scale bars represent 10 mm. Shown are the fluorescence signals of a-bungarotoxin staining (E1) and of YFP emitted from RIa-EPAC and (E2). (F1 and F2) YFP-fluorescence signals upon two-photon excitation at 820 nm of RIa-EPAC at different time points before and after injection of the cAMP agonist, forskolin (FRSK). (F3 and F4) Pseudocolored ratiometric images showing the ratio of CFP/YFP emission signals upon two-photon excitation at 820 nm. The scale bar beneath F4 indicates absolute ratio values. Note that only after application of FRSK a clear increase of the CFP/YFP ratio can be observed. (G) Graphical representation of the mean CFP/YFP ratio changes SEM (n ¼ 13 synapses) upon FRSK application.
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acquired in single-photon mode (synapse, Fig. 19.3E1 PKA I marker, Fig. 19.3E2), which was then analyzed by multiphoton imaging to determine its cAMP signals (Fig. 19.3F and G). Figure 19.3F1 and F2 show the yellow fluorescent protein (YFP) emission images upon multiphoton excitation at 820 nm at different time points before and after cAMP agonist application. Figure 19.3F3 and F4 depict the corresponding ratiometric images, which were created as described in Section 2.4. As already mentioned, combinations of this and other imaging modes allow to quantify a set of parameters in one and the same sample and therefore to correlate second messenger signals to their direct or indirect outputs. Due to the wide range of applications of multimodal imaging, we cannot discuss the requirements for every type of imaging. Therefore, in this chapter, we give only general remarks as to the acquisition of FRETbased second messenger signals. First, make sure that your additional dyes do not interfere with the FRET-fluorescence signals. We have made good experience with infrared stains, such as AlexaFluor 647 in combination with cyan fluorescent protein (CFP)–YFP FRET pairs (Fig. 19.3). The use of optical parameter oscillators (OPOs), which broaden the tunable multiphoton spectrum to about 1300 nm (current commercially available multiphoton lasers for microscopy stop at about 1000 nm wavelength), might extend the use of multimodal microscopy to additional dye combinations. Second, different types of FRET imaging can be pursued, but not all of them can be easily used for quantitative assessments. It is fair to say that FRET– fluorescence-lifetime imaging microscopy (FLIM) applications might be the most quantitative, but they are still pretty slow and, at least to our knowledge, they have not been described in living animals so far. Among the other available methods to detect FRET, we use dynamic ratiometric FRET imaging, which relies on increased FRET acceptor (e.g., YFP) emission at the expense of donor (e.g., CFP) emission upon donor excitation, when FRET efficiency is rising. Although it is not as quantitative as FRET–FLIM, simple division of data from simultaneously acquired image pairs (donor and acceptor emission) leads to quick and relatively movement-resistant insights into the dynamism of second messenger signals. In the case of CFP as a donor, multiphoton excitation at 820 nm has proven to bear very low crosstalk with YFP acceptors. Excitation spectra with samples expressing only one of the two FRET pair dyes permit identifying the wavelength where the amount of crosstalk is minimal. FRET image pairs should be taken at 12-bit image depth and sensors should be set to linearly respond to a wide range of incoming light. It is instrumental to work in the optimal gain range of your detectors and to keep these values constant over all your experiments. Also avoid using detector-offset values deviating strongly from zero, because this will lead to an increasingly nonlinear data acquisition. Finally, when using very different excitation and emission wavelengths, the inherent optical properties of lasers and lenses render a
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perfect alignment in x–y as well as in z very difficult. In other words, sequential use of short- and long-wavelength light might excite your sample at distinct planes. Additionally, the emission signals might undergo a differential movement along the optical path. These parameters should be checked for each new optical component, for example, using fluorescent beads. 2.3.1. Instrumentation and disposables needed 2.3.1.1. Instruments Upright microscope equipped with dual capabilities for measuring in singleand multiphoton mode Water immersion objectives for direct immersion with optimal correction for spherical and chromatic aberrations as well as high numerical aperture Custom-made support for mounting the animal onto the microscope object table Straight 9-cm surgical scissors Dumont #7 curved shanks forceps (11.5 cm) with and without serrations Narrow pattern curved forceps (12 cm) Hartman straight hemostat (10 cm) for fixing distal tendon to metal support Additional hardware (e.g., electrodes, force transducer) depends on the type of parameters to be analyzed 2.3.1.2. Solutions
Physiological solution to moisten the muscle during the preparation Anesthetics: Zoletil 100 (Virbac) and Rompun (2% xylazine, Bayer) Dye solutions in sterile physiological solution (if applicable) ViscOphtal (Winzer Pharma), artificial tear gel for objective immersion 2.3.1.3. Disposables
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2.4. Data extraction Data analysis again widely depends on which parameters need to be detected. For data analysis, we use ImageJ, a freeware, which was generated by the NIH (Wayne Rasband, NIH, Bethesda, USA) and now uses a broad base of customers, which participate in programming (http://rsb.info.nih.gov/ij/). We describe here the main features to analyze the dynamic FRET-based second messenger signals. First, subtract background from your images. In ImageJ, this can be done using the Process/Math/Subtract prompt. In most confocal or multiphoton applications, background is relatively low and stable and therefore a generic subtraction of a mean of background values measured outside the
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transfected area is mostly sufficient. Strong variations in background levels are often due to major changes in focus or due to alterations in tissue composition, such as upon influx of blood or lymph into the observed region. In such cases, a reliable background subtraction is difficult to achieve and data must be handled with extreme caution. If the origin of such deviations can be identified, try to eliminate or reduce it. After background subtraction, regions of interest need to be determined. Ideally, the signal of interest is so strongly above background that simply the entire signal above background can be used for data analysis. Alternatively, additional operations have to follow. We usually generate masks on the composites of the background-subtracted donor and acceptor images. For the composite, use the Add function under Process/ImageCalculator. Then, you may threshold pixels to be analyzed by using the Image/Adjust/ Threshold command. The Apply threshold command will lead to the creation of a mask, which, after setting minimum and maximum to 0 and 1, respectively, can be multiplied with the background-subtracted donor and acceptor images. The minimum–maximum setting can be achieved by dividing the mask images by 255. Multiplication with the donor and acceptor images can be done using the multiply function under Process/ImageCalculator. The resulting images can be used to create a ratiometric image file using the CalculatorPlus plugin. This allows dividing donor and acceptor images with an additional constant to avoid clipping of decimal places of ratio values (note that output values are always integers). For calculating the CFP/YFP ratio in the CalculatorPlus function Divide, i2 ¼ (i1/i2) k1 þ k2, we use donor and acceptor images as i1 and i2, respectively. This is good in sensors, where increase in a second messenger leads to reduced FRET, such as in EPAC sensors for cAMP. The inverse is the case with cameleon probes, where donor and acceptor images make i2 and i1, respectively. In all cases, a k1 value of 5000 avoids clipping of minima and maxima, and k2 is zero. Using such ratiometric images, the dynamics of second messenger signals can either be globally or locally determined. For global analyses, it is sufficient to read out the entire ratiometric images with a threshold higher than zero (in order to get rid of the background pixels). For local analyses, regions of interest might need to be adapted according to specific needs. For all these calculations good standard PCs are sufficient.
3. Adaptations to Tissues Other Than Skeletal Muscle 3.1. Isolated heart Single-photon confocal microscopy has extensively been used to study second messenger signaling in isolated cardiac cells. Use of a vast number of chemical- and protein-based reporters in single- and multimodal imaging experiments has allowed simultaneous interrogation of intracellular messengers including Ca2 þ, cAMP, or biophysical parameters such as
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membrane potential. The functional properties of cardiomyocytes, however, are heavily influenced by a number of factors including specific interactions with neighboring cardiomyocytes, fibroblasts, and the extracellular matrix, none of which are preserved in isolated cardiomyocytes in vitro. The ability to measure variations in intracellular second messengers at subcellular level within the intact tissue would thus provide key information to understand cellular physiology and pathophysiology of the heart. However, a relatively small number of studies have been conducted addressing second messenger imaging in rodent cardiac tissue (i.e., papillary muscles; Wier et al., 1997) or intact hearts, using either surface imaging (Minamikawa et al., 1997) or deep tissue imaging with multiphoton microscopy (Aistrup et al., 2006; Rubart et al., 2003). These models are especially relevant to infer the mechanism of cardiovascular diseases like heart failure, featuring extracellular matrix remodeling and fibroblast growth, or arrhythmias, for example, those resulting from defective cell to cell communication, whereby behavior of cardiomyocytes in their tissue environment may differ substantially from that of the isolated cell. Here, we illustrate an adaptation of the method described in detail above for skeletal muscle imaging to the use in the isolated, Langendorff-perfused mouse heart. We will focus on the protocol for multiphoton imaging with the Ca2 þ indicator fluo-4, although with slight variations, the same setup can be utilized with FRET-based sensors to detect intracellular levels of other second messengers. 3.1.1. Probe introduction into mouse hearts A number of different approaches can be used to obtain expression of fluorescent indicators in the mouse heart. These can be summarized as follows: transgenic expression, viral transduction either via systemic or local intramyocardial injection, and dye loading in the isolated heart. 3.1.2. Heart preparation Ten minutes after heparinization (heparin, 200 IU/kg, i.p.), the heart of a 2- to 3-month-old mouse is excised and the aorta cannulated with a blunt 18 G needle (Fig. 19.4A and B). Retrograde coronary perfusion of oxygenated Tyrode solution is achieved at room temperature, at a constant pressure of 70 mmHg, and carried on for 15–20 min to equilibrate. At this point, if dye loading is needed, a mixture of Fluo-4AM (10 mM)/ Pluronic F127 (10 mM) and sulfinpyrazone (SP, 10 mM) is added to the solution and perfused for 20 min, after which the heart is perfused with SP containing dye-free Tyrode solution for 30 min to allow de-esterification of the AM dye.
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Figure 19.4 Summary of the isolated heart Ca2 þ imaging experiment. (A) Schematic of isolated heart imaging. (B, C) Intact isolated mouse heart imaging involves three consecutive steps: (i) heart harvesting, aortic cannulation, and retrograde perfusion setup (B); (ii) dye loading by perfusion recirculation; and (iii) multiphoton imaging during electrical pacing (C). (D) Images recorded from subepicardial fibers of a fluo3AM-loaded mouse heart, during electrical pacing via the aortic cannula. Images were acquired at a rate of 8 frames/s during a systolic Ca2 þ transient and represented in intensity-weighed pseudocolor using a custom-made software routine operating in Matlab (Vimmaging, F. Mammano and C. Ciobutaru, Padova, Italy).
3.1.3. Heart imaging Tyrode solution perfusion is continued while the heart is mounted horizontally on a chamber made of an agarose cast (low melting point agarose, 4% in Tyrode; Fig. 19.4C), which is used to keep the heart in position under the microscope objective (Fig. 19.4A). To minimize movement, we use a combination of the contraction inhibitors butanedione monoxime (BDM, 10 mM) and blebbistatin (100 nM). A bipolar pulse generator (Ionoptix, USA) connected to the cannula and to a ground electrode embedded in the agarose cast is used to pace the heart. During imaging (Fig. 19.4C), a drop of tear gel is placed on the epicardial surface. When
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imaged with multiphoton microscopy, the first 8–10 subepicardial cardiomyocyte layers on the left ventricular wall appear oriented along the long axis of the heart and can be efficiently visualized (Fig. 19.4D). 3.1.4. Instrumentation and disposables needed 3.1.4.1. Instruments Upright microscope equipped with dual capabilities for measuring in singleand multiphoton mode Water immersion objectives for direct immersion with optimal correction for spherical and chromatic aberrations as well as high numerical aperture Straight 9-cm surgical scissors Dumont #7 curved shanks forceps (11.5 cm) Narrow pattern curved forceps (12 cm) Bipolar pulse isolator (i.e., MyoPacer Cell Stimulator, Ionoptix, USA) 22G Aortic Cannula for mouse heart (Cat No. SP3787, ADI Instruments, UK) 3.1.4.2. Solutions
Heart perfusion solution: HEPES-buffered Tyrode solution (HE-Tyrode), 100% O2 gassed Dye loading solution (if applicable): HE-Tyrode, Fluo-4AM (10 mM)/Pluronic F127 (10 mM), SP Dye washout solution: HE-Tyrode, SP Imaging solution: HE-Tyrode, BDM (10 mM), ()-Blebbistatin (100 nM, Tocris, MI, USA) Heparin, 200 IU/kg in PS Low melting point agarose ViscOphtal (Winzer Pharma), artificial tear gel for objective immersion
3.2. Brain imaging—A brief teaser Understanding signaling of neuronal circuits requires the monitoring of electric activities. Therefore, action potentials (APs) emitted by neurons are one of the most important functional parameters. APs have fast dynamics in the millisecond time scale and allow specific communication with other cells via spatio-temporal patterns. Because of their unparalleled temporal resolution, one would ideally utilize electrophysiological methods for intraand intercellular recordings within local networks and during behavior. However, the experimenter is faced with the challenge to apply them to behaving animals or to more than one cell at a time. An alternative approach and a fundamental breakthrough in general has been the use of multiphoton microscopy in combination with fluorescent probes sensitive to cellular processes, and appropriate for monitoring of synaptic activity (Dreosti and Lagnado, 2011; Grewe and Helmchen, 2009; Helmchen and Denk, 2005).
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Unfortunately, genetically encoded voltage-sensitive dyes, made by either fusing FPs to the Shaker Kþ channel (FlaSh), sodium channels, or built around voltage-sensitive phosphatases, still produce signals that are too small and slow to be used at the level of single cells. This limitation of low signalto-noise ratio needs to be overcome in the future by the development of new high-performance voltage sensors (Mutoh et al., 2011). However, in neurons, APs usually result in a transient increase in intracellular Ca2 þ concentrations. Thus, Ca2 þ imaging is currently the most promising approach for neuronal activity sampling in all cells within a region of hundreds of micrometers across, and at cellular resolution within local neuronal circuits (Grewe et al., 2010). Some studies have used bulk loading of membrane-permeant chemical Ca2 þ dyes, such as fluo-3, Oregon Green 488, or rhod-2. They can be introduced into thousands of cells through a small skull opening and pressure ejection (Garaschuk et al., 2006). This technique, termed multicell bolus loading, results in a rather uniform staining of neurons within the targeted area and also proved to be well suited for pharmacological studies in awake animals, for example, after local administration of drugs. However, since Ca2 þ transients form with a delay of a few milliseconds after the AP, and because Ca2 þ transients decay much slower than APs, Ca2 þ imaging has some limitation in the precise decoding of individual APs at high firing rates. In addition, bolus loading of chemical dyes lacks discrete indicator labeling of specific cell types. Recent studies have addressed these challenges. First, Grewe et al. (2010) greatly improved the temporal resolution of AP-evoked Ca2 þ transients for in vivo imaging in mouse neocortex down to 300 mm using random access scanning with acousto-optic deflectors (AOD). Notably, the AOD-based two-photon microscope allowed sampling rates of up to 500 Hz and determined spike times with near millisecond precision and only 5–15 ms confidence intervals. Although in vivo measurements were collected in two dimensions, this approach should be further extended using z-dimension scanning. Second, Drobizhev et al. (2011) characterized the two-photon absorption (2PA) spectra of 48 FPs, from enhanced BFP- and CFP-series, the “fruit” FPs to far-red variants, such as mRaspberry, mKate2, tandem dimer (td) Katushka2, or eqFP670. This approach will lead to the development of brighter and faster mutants for multiphoton applications. However, there are other factors of concern—such as photostability, photoswitching efficiency, and cellular expression rate. While some FP constructs like YC-cameleons under the control of constitutively active promoters proved to be critical (Nagai et al., 2004), similar indicators using troponin-C instead of calmodulin as a Ca2 þ sensing domain placed under the control of an inducible promoter were suitable to detect fast and robust Ca2 þ transients in neurons of transgenic mice (Hasan et al., 2004). However, the generation of transgenic mice is time consuming and often the overall success rate is moderate. A faster alternative is the use of lenti- or adenovirus-based systems for gene delivery to postmitotic cells. They offer several advantages,
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as stereotactic injection makes it easy to deliver recombinant probes locally and to control expression rate in a defined time window (Lutcke et al., 2010; Osten et al., 2006). Further, fluorescent labeling of a small population of neurons, in an otherwise unlabeled tissue, offers a high signal-to-noise ratio.
4. Concluding Remarks The monitoring of second messenger signaling in living animals and intact tissues is now an established method. In contrary to single cell-based assays, however, the modes of targeting the sensor into the tissue of interest and the challenges encountered during the imaging process greatly vary from tissue to tissue, and thus in each case, the methodological setup needs careful adaptation. This chapter elaborated on these aspects in three types of tissue, that is, skeletal muscle, heart muscle, and neuronal tissue to exemplify some of the most immediate practical challenges. We are aware that the concerns are likely to be different for other organs. In parallel with continuous refinement of the experimental procedures and with the generation of fluorescent probes more suited to multiphoton excitation, technical developments are rapidly leading to faster and more powerful microscopes. This combination is expected to enable in the near future increasingly better spatial and temporal resolution in the detection of biologically and pathologically relevant events in intact tissues and organs.
ACKNOWLEDGMENTS We apologize to all authors whose work could not be included due to space limitations. We are grateful to Anika Wagner for excellent technical assistance. The work of R. R. is supported by the Helmholtz Association, the Deutsche Forschungsgemeinschaft, and the Association Franc¸aise contre les Myopathies. Financial support from the Ministry of Science, Research and Art (BadenWu¨rttemberg—“Innovative Projekte”) to M. H. is gratefully acknowledged. M. M. is supported by the EU Seventh Framework Program FP7/2007-2013 (HEALTH-F2-2009-241526, EUTrigTreat).
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Dona, M., Sandri, M., Rossini, K., Dell’Aica, I., Podhorska-Okolow, M., and Carraro, U. (2003). Functional in vivo gene transfer into the myofibers of adult skeletal muscle. Biochem. Biophys. Res. Commun. 312, 1132–1138. Dreosti, E., and Lagnado, L. (2011). Optical reporters of synaptic activity in neural circuits. Exp. Physiol. 96, 4–12. Drobizhev, M., Makarov, N. S., Tillo, S. E., Hughes, T. E., and Rebane, A. (2011). Two-photon absorption properties of fluorescent proteins. Nat. Methods 8, 393–399. Garaschuk, O., Milos, R. I., and Konnerth, A. (2006). Targeted bulk-loading of fluorescent indicators for two-photon brain imaging in vivo. Nat. Protoc. 1, 380–386. Grewe, B. F., and Helmchen, F. (2009). Optical probing of neuronal ensemble activity. Curr. Opin. Neurobiol. 19, 520–529. Grewe, B. F., Langer, D., Kasper, H., Kampa, B. M., and Helmchen, F. (2010). High-speed in vivo calcium imaging reveals neuronal network activity with near-millisecond precision. Nat. Methods 7, 399–405. Hasan, M. T., Friedrich, R. W., Euler, T., Larkum, M. E., Giese, G., Both, M., Duebel, J., Waters, J., Bujard, H., Griesbeck, O., Tsien, R. Y., Nagai, T., et al. (2004). Functional fluorescent Ca2 þ indicator proteins in transgenic mice under TET control. PLoS Biol. 2, e163. Helmchen, F., and Denk, W. (2005). Deep tissue two-photon microscopy. Nat. Methods 2, 932–940. Lutcke, H., Murayama, M., Hahn, T., Margolis, D. J., Astori, S., Zum Alten Borgloh, S. M., Gobel, W., Yang, Y., Tang, W., Kugler, S., Sprengel, R., Nagai, T., et al. (2010). Optical recording of neuronal activity with a genetically-encoded calcium indicator in anesthetized and freely moving mice. Front. Neural Circuits 4, 9. Minamikawa, T., Cody, S. H., and Williams, D. A. (1997). In situ visualization of spontaneous calcium waves within perfused whole rat heart by confocal imaging. Am. J. Physiol. 272, H236–H243. Mutoh, H., Perron, A., Akemann, W., Iwamoto, Y., and Knopfel, T. (2011). Optogenetic monitoring of membrane potentials. Exp. Physiol. 96, 13–18. Nagai, T., Yamada, S., Tominaga, T., Ichikawa, M., and Miyawaki, A. (2004). Expanded dynamic range of fluorescent indicators for Ca(2þ) by circularly permuted yellow fluorescent proteins. Proc. Natl. Acad. Sci. USA 101, 10554–10559. Osten, P., Dittgen, T., and Licznerski, P. (2006). Lentivirus-based genetic manipulations in neurons in vivo. In “The Dynamic Synapse: Molecular Methods in Ionotropic Receptor Biology,” (J. T. Kittler and S. J. Moss eds.). CRC Press, Boca Raton, FL. Pozzan, T., Mongillo, M., and Rudolf, R. (2003). The Theodore Bucher lecture. Investigating signal transduction with genetically encoded fluorescent probes. Eur. J. Biochem. 270, 2343–2352. Ro¨der, I. V., Petersen, Y., Choi, K. R., Witzemann, V., Hammer, J. A., 3rd, and Rudolf, R. (2008). Role of Myosin Va in the plasticity of the vertebrate neuromuscular junction in vivo. PLoS One 3, e3871. Ro¨der, I. V., Lissandron, V., Martin, J., Petersen, Y., Di Benedetto, G., Zaccolo, M., and Rudolf, R. (2009). PKA microdomain organisation and cAMP handling in healthy and dystrophic muscle in vivo. Cell. Signal. 21, 819–826. Ro¨der, I. V., Choi, K. R., Reischl, M., Petersen, Y., Diefenbacher, M. E., Zaccolo, M., Pozzan, T., and Rudolf, R. (2010). Myosin Va cooperates with PKA RIalpha to mediate maintenance of the endplate in vivo. Proc. Natl. Acad. Sci. USA 107, 2031–2036. Romanello, V., Guadagnin, E., Gomes, L., Ro¨der, I., Sandri, C., Petersen, Y., Milan, G., Masiero, E., Del Piccolo, P., Foretz, M., Scorrano, L., Rudolf, R., et al. (2010). Mitochondrial fission and remodelling contributes to muscle atrophy. EMBO J. 29, 1774–1785.
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Rubart, M., Wang, E., Dunn, K. W., and Field, L. J. (2003). Two-photon molecular excitation imaging of Ca2 þ transients in Langendorff-perfused mouse hearts. Am. J. Physiol. Cell Physiol. 284, C1654–C1668. Rudolf, R., Mongillo, M., Magalhaes, P. J., and Pozzan, T. (2004). In vivo monitoring of Ca2 þ uptake into mitochondria of mouse skeletal muscle during contraction. J. Cell Biol. 166, 527–536. Rudolf, R., Magalhaes, P. J., and Pozzan, T. (2006). Direct in vivo monitoring of sarcoplasmic reticulum Ca2 þ and cytosolic cAMP dynamics in mouse skeletal muscle. J. Cell Biol. 173, 187–193. Tothova, J., Blaauw, B., Pallafacchina, G., Rudolf, R., Argentini, C., Reggiani, C., and Schiaffino, S. (2006). NFATc1 nucleocytoplasmic shuttling is controlled by nerve activity in skeletal muscle. J. Cell Sci. 119, 1604–1611. Valkova, C., Albrizio, M., Ro¨der, I. V., Schwake, M., Betto, R., Rudolf, R., and Kaether, C. (2011). Sorting receptor Rer1 controls surface expression of muscle acetylcholine receptors by ER retention of unassembled alpha-subunits. Proc. Natl. Acad. Sci. USA 108, 621–625. Whitaker, M. (2010). Genetically encoded probes for measurement of intracellular calcium. Methods Cell Biol. 99, 153–182. Wier, W. G., ter Keurs, H. E., Marban, E., Gao, W. D., and Balke, C. W. (1997). Ca2 þ ‘sparks’ and waves in intact ventricular muscle resolved by confocal imaging. Circ. Res. 81, 462–469. Zaccolo, M., and Pozzan, T. (2003). cAMP and Ca(2 þ) interplay: A matter of oscillation patterns. Trends Neurosci. 26, 53–55. Zaccolo, M., Magalhaes, P., and Pozzan, T. (2002). Compartmentalisation of cAMP and Ca (2 þ) signals. Curr. Opin. Cell Biol. 14, 160–166. Zhang, J., Campbell, R. E., Ting, A. Y., and Tsien, R. Y. (2002). Creating new fluorescent probes for cell biology. Nat. Rev. Mol. Cell Biol. 3, 906–918.
N I N E T E E N
Investigating Second Messenger Signaling In Vivo ¨diger Rudolf,* Mathias Hafner,† and Marco Mongillo‡ Ru Contents 1. Introduction 2. Multimodal Imaging in Living Mouse Skeletal Muscle 2.1. Ad hoc probe introduction: Injection, electroporation 2.2. Preparation of the mouse for imaging: Time for add-ons 2.3. In vivo imaging of mouse skeletal muscle: Space limitations 2.4. Data extraction 3. Adaptations to Tissues Other Than Skeletal Muscle 3.1. Isolated heart 3.2. Brain imaging—A brief teaser 4. Concluding Remarks Acknowledgments References
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Abstract All known second messengers are small molecules without complex structural features. However, each of them can mediate different, very specific cellular responses upon arrival of distinct extracellular cues in one and the same cell. From this follows the question of how signal specificity is achieved in space and time. Recent work showed that three factors play major roles in determining signal specificity: intensity, pattern, and compartmentalization of signals. Thus, for understanding the signaling in any specific context, generic information on the involvement of second messenger pathway(s) is insufficient and only the precise, time- and space-resolved monitoring of these processes will lead to meaningful insights. Even more demanding, many second messenger-based signaling events can only occur using tissue-specific morphological arrangements (e.g., striated organization of muscle and synaptic specializations in neurons), making the visualization of intact tissue mandatory. Finally, to * Institut fu¨r Toxikologie und Genetik, Karlsruhe Institute of Technology, Hermann-von-Helmholtz-Platz 1, Eggenstein-Leopoldshafen, Germany Institut fu¨r Medizintechnologie der Universita¨t Heidelberg und Hochschule Mannheim, Mannheim, Germany { Venetian Institute of Molecular Medicine, Via Orus 2, Padua, Italy {
Methods in Enzymology, Volume 505 ISSN 0076-6879, DOI: 10.1016/B978-0-12-388448-0.00027-9
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2012 Elsevier Inc. All rights reserved.
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appreciate the physiological relevance of second messenger signals, multiparametric readouts are generally needed. This chapter illustrates modes of multiparametric analysis of signaling in live mouse skeletal muscle and briefly discusses possible applications of the technique in other tissues including heart and brain.
1. Introduction Second messengers mediate an enormous spectrum of cellular responses to external stimuli, ranging from the regulation of cell proliferation and metabolism to cell death. Notably, different signals might arise within one cell at the same time, rendering it absolutely necessary for the cell to achieve signal specificity (Zaccolo and Pozzan, 2003; Zaccolo et al., 2002). Clearly, a vast proportion of this specificity is due to amplitude, pattern (e.g., oscillations), and subcellular compartmentalization of signals. In particular, the latter is completely cell-type dependent and can usually not be reconstituted in vitro in a reliable manner. Therefore, the physiological roles of second messenger signaling can often only be understood when studied in the living tissue or in the intact organism. Since the discovery of green fluorescent protein (GFP), numerous fluorescent, genetically encoded sensors able to monitor second messengers in living cells were created and new ones are continuously developed (Pozzan et al., 2003; Whitaker, 2010; Zhang et al., 2002). In combination with adequate approaches, genetically modified organisms, and molecular or electronic biosensors measuring additional biophysical parameters, the use of GFP-based sensors can now deliver invaluable insights into many second messenger-dependent physiological processes in intact tissue. In most cases, probes for second messenger dynamics exploit Fo¨rster resonance energy transfer (FRET), usually between cyan and yellow fluorescent proteins (FPs), to transform conformational changes in the sensor molecule into measurable changes in their fluorescence spectra. The investigation of such probes in living samples has been enabled by multiphoton imaging, which allows excitation of short-wavelength dyes deep in tissue. However, the merit of tissue penetration of multiphoton light is dampened by its broad excitation range, which makes the discrimination of more than two fluorescent probes difficult. However, since dyes with longer wavelengths can efficiently be visualized even with standard confocal microscopy deep in tissue, multimodal imaging, using multiphoton together with single-photon confocal microscopy is useful to combine the visualization of second messenger signals with other readouts. Transgenic techniques have rendered possible the introduction of genetically encoded probes into mice and a few other model organisms.
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However, at least in mammals, these methods cannot cope with the speed of novel sensors development and the progress in biomedical research. Therefore, faster approaches to introduce molecular sensors into target tissues are needed. While for some organs viral vectors might be ideal, electroporation-mediated gene transfer has proven to be an efficient means in skeletal muscle. This chapter first describes the steps from genetic manipulation of skeletal muscle to the multimodal imaging and data analysis of second messenger signaling and other factors in this tissue in living mice. Then, we introduce adaptations of this paradigm to other tissues, namely, cardiac muscle and neuronal tissue.
2. Multimodal Imaging in Living Mouse Skeletal Muscle Second messenger monitoring alone can be used to investigate the effect of ligands or trigger stimuli on a selected second messenger pathway. However, to understand the physiological meaning of such a signal, second messenger monitoring needs to be coupled to additional readouts. This can be done by combining in vivo imaging with electronic biosensors, such as force transducers (Allen et al., 2011) or patch clamp devices. Alternatively, if fluorescent long-wavelength markers are available, these can be analyzed by using single-photon confocal microscopy in combination with second messenger analysis of FRET-based probes in the multiphoton mode (Ro¨der et al., 2009, 2010). Additional combinations, for example, with second harmonic generation imaging can be envisaged. The ability of multimodal imaging to interrogate simultaneously intracellular signals and functional outputs makes a clear advantage over the single modal analysis. In the following paragraphs, the steps from probe introduction to multimodal imaging and data analysis are exemplified for mouse hindlimb muscle.
2.1. Ad hoc probe introduction: Injection, electroporation Due to their accessibility, muscles of the mouse distal extremities are particularly amenable to probe injection and electroporation (Figs. 19.1A and B and 19.2A–G). On the one hand, injected probes might be cell-permeant or cell-impermeant, such as tetramethyl rhodamine methyl ester (TMRM) (a mitochondrial membrane potential marker; Romanello et al., 2010) or a-bungarotoxin coupled to chromophores (marking AChRs in neuromuscular junctions; Ro¨der et al., 2008, 2010), respectively. On the other hand, electroporation (Figs. 19.1B and 19.2A–G) is a quick and efficient method to introduce cDNAs for expressing genetically encoded fluorescent probes or
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A Injection of cDNA and/or dyes
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Figure 19.1 Schematic of in vivo-imaging procedure. Typical experiments involving live imaging of mouse skeletal muscle are divided into at least three phases. First, muscles are prepared by probe and/or cDNA injection (A). Second, in many cases, muscle transfection is used to incorporate sensor-encoding cDNAs and/or muscle function-modifying constructs (B). Third, a couple of days later, in vivo imaging is performed, often using multiple excitation and emission modes to gather multifactorial readout data (C). In some cases, additional treatments (e.g., pharmacological or surgical interventions) are adopted (not shown).
other genes or siRNAs, which modulate any tissue function of interest. As shown previously, transfected cDNAs remain actively transcribed and translated for up to 4 weeks after electroporation in skeletal muscle (Dona et al., 2003). This is most likely due to the postmitotic status of adult skeletal muscle. Although very high levels of up to 80% of transfected fibers might be reached
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Figure 19.2 Setups and procedures for transfection and in vivo imaging of mouse skeletal muscle. (A–G) Setup and procedure of muscle transfection. A setup composed of electroporator, electrodes, heated operation platform, and surgical instruments is used (A). Electroporation involves shaving of the lower hindlimb (B), a longitudinal cut to get access to the hindlimb muscles (C), insertion of a spatula electrode (D), injection of cDNA (E), application of the second electrode and electropulsing (F), and closure of the wound by surgical stitches (G). (H–N) Preparation for in vivo imaging. Ten days after electroporation, the wound should be completely healed (H) and the muscle exhibit a pale rosy color (I). After removing the skin above the electroporated muscle (here: tibialis anterior), the transparent epimysium covering the muscle is eliminated (J). Using forceps and scissors, the distal tendon is cut (K, L) and the mouse transferred to a custom-made table for microscopy (M). Using a hemostatic clamp and plastic and cork supports, the distal tendon is then fixed over plastic tubing (N). (O–Q) Transfer to microscope and in vivo imaging. A confocal microscope equipped with an upright stand, standard and two-photon lasers, and water immersion objectives adapted for use without coverglass is used (O) and the custom support is mounted on the microscope object table (P, Q).
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for some constructs, expression of heterologous proteins being targeted to specific sites or organelles, such as the synaptic membrane, mitochondria, or the sarcoplasmic reticulum can be much more cumbersome. This is particularly true for transmembrane proteins of the synapse, since the secretory pathway mediating the delivery of synaptic proteins is scarcely developed in skeletal muscle. For two main reasons, we usually wait a couple of days (10) before imaging transfected muscles. First, a mild, unavoidable inflammation typically occurring at the site of electroporation is completely healed by then. Second, many proteins in skeletal muscle exhibit low turnover rates, such as the AChR, which has a half-life of >10 days. It is also critical to have always the same, experienced persons performing the electroporation, because this strongly reduces damaging of muscle, which is almost zero in the optimal case, but can be problematic upon inadequate performance. It is worth noting that electroporation efficiency might vary from mouse strain to mouse strain and it is particularly low in muscles with a lot of fat and connective tissue, such as in dystrophic animals. Of the different protocols available for muscle transfection, we prefer to use the method from Dona et al. (2003), where the transfected muscle (e.g., tibialis anterior (TA)) is exposed by simple surgery (Figs. 19.1A, B and 19.2A–G). This is ideal for local and muscle-specific transfection and allows transfecting either the whole muscle (by injecting at different sites) or only superficial fiber layers (sufficient for imaging). For transfecting lower hindlimb muscles, such as TA or extensor digitorum longus (EDL), we use spatula electrodes of 1 mm thickness and 4 mm width (Fig. 19.2D). These are connected to a BTX ECM830 square pulse generator (Fig. 19.2A). After orienting the anesthetized mouse on the side to be transfected, the lower hindlimb is stretched over a pillow and crepe tape is used to fix the foot (Fig. 19.2B). After shaving the leg, a longitudinal cut of 6 mm length opens the midline of the lower hindlimb exposing a view on the whitish border between anterior and posterior muscles (Fig. 19.2C). The shiny fascia (see Fig. 19.2J) enclosing this whole arrangement is then opened by a second longitudinal cut. This allows separating anterior from posterior muscles and seeing the tibia. Now, a closed pair of small surgical scissors penetrates between the tendon of the TA and the tibia. Careful opening of the scissors creates a hole through which the electrode can be inserted (Fig. 19.2D). After fixing the electrode on the pillow with tape, a Hamilton syringe is used to slowly inject up to 20 ml of cDNA in physiological solution (less is better, Fig. 19.2E). Depending on the construct, 0.5–20 mg of cDNA might be used. Make sure that no liquid is leaving the injection site, because this usually results in low transfection efficiency. If necessary, additional, cell-impermeant labels should be injected subsequently. Then, carefully place the second electrode on top of the muscle (Fig. 19.2F) and apply pulses in the following pattern: five pulses with 5 ms duration each are given at an interval of 200 ms between pulses. Voltage is
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typically 50 V/cm of muscle. Make sure to adapt the voltage to muscle thickness. Then, carefully remove both electrodes and close the skin with two to four surgical stitches (Fig. 19.2G), followed by sterilizing the wound. Experienced operators perform a transfection in a few minutes, and animals immediately walk, jump, and behave normally after complete recovery from anesthesia. To help fast recovery, let the animals wake up on a heating plate at 37 C. Animals, which do not behave normally few hours after transfection, suffer from pain or infection and should additionally be treated with an antibiotic and analgesic. 2.1.1. Instrumentation and disposables needed 2.1.1.1. Instruments BTX ECM830 square wave pulse generator Two spatula electrodes and cables Heating plate set to 37 C Hamilton syringe (20 or 100 ml) Straight 9-cm surgical scissors Dumont #7 curved shanks forceps (11.5 cm) with and without serrations Narrow pattern curved forceps (12 cm) Surgical needles (size 2) Needle holder (12.5 cm) Surgical silk (size 5/0) Surgery plate for small animals Custom-made pillow to sustain leg for transfection 2.1.1.2. Solutions
Physiological solution to moisten the muscle during the transfection procedure Anesthetics cDNA and other dye solutions in sterile physiological solution 2.1.1.3. Disposables
Crepe tape
2.2. Preparation of the mouse for imaging: Time for add-ons A few days after transfection, the surgical wound should be completely healed (Fig. 19.2H), and when opening the skin of the anesthetized mouse for imaging, the transfected muscle should exhibit a nicely smooth surface and a pale rosy color (Fig. 19.2I). Any sign of inflammation or hematoma indicates severe muscle damage and is strongly counterindicating the use of
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such a specimen for imaging. Healthy muscles can be further processed for in vivo imaging. Depending on the experiment, muscles can again be injected with fluorescent dyes (Ro¨der et al., 2008, 2010; Romanello et al., 2010; Valkova et al., 2011), or they can be attached to force transducers (Allen et al., 2011), muscle electrodes (Rudolf et al., 2004, 2006; Tothova et al., 2006), or other electronic devices. Anesthesia might be adapted to specific needs, but for imaging, we have made the best experience using intraperitoneal injection of a mixture of Zoletil 100 and Rompun. Compared to other anesthetics, there is much less spontaneous movement of muscles, and anesthesia of the animal is deep and reliable for a time period of about 2 h. Also, orienting and fixing the observed muscle properly is crucial. We detach the distal tendon of the TA (Fig. 19.2K and L) and fix it either with a surgical hemostat or hook it with surgical silk to a force transducer in an angle of 70 –80 from the tibia and horizontal to the surface of the support (Fig. 19.2M and N). This makes the muscle more rigid for imaging and further reduces breathing-induced muscle movements. Before detaching the tendon, carefully remove the fascia directly surrounding the muscle (epimysium, Fig. 19.2J). If electroporation was carried out properly, this layer should still be intact, and after a careful longitudinal cut from the distal tendon to the knee, the two halves of the epimysium can be flipped to both sides of the muscle. Make attention to moisten the muscle frequently with physiological solution. In case of muscle force measurements, the tibia has to be properly fixed to the support using hooks. For pure imaging, the use of crepe tape placed over different points of the leg and the body is sufficient. It is ideal to use a custommade metal support with threads at different positions (Fig. 19.2M) to fix the table after mounting of the animal on the microscope table and to position further tools, such as micromanipulators, force transducers, etc. 2.2.1. Instrumentation and disposables needed 2.2.1.1. Instruments Custom-made support for mounting the animal onto the microscope object table Straight 9-cm surgical scissors Dumont #7 curved shanks forceps (11.5 cm) with and without serrations Narrow pattern curved forceps (12 cm) Hartman straight hemostat (10 cm) for fixing distal tendon to metal support Surgery plate for small animals 2.2.1.2. Solutions
Physiological solution to moisten the muscle during the transfection procedure Anesthetics: Zoletil 100 (Virbac) and Rompun (2% xylazine, Bayer) Dye solutions in sterile physiological solution (if applicable)
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2.2.1.3. Disposables
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2.3. In vivo imaging of mouse skeletal muscle: Space limitations Once mounted on the metal support, the mouse is transferred to the microscope for imaging. Use an upright microscope, which permits working both in single-photon confocal as well as multiphoton mode (Fig. 19.2O). The object table of the microscope should permit application of your custom-made support carrying the prepared mouse and electronic z-plane motion (Fig. 19.2P). At its lowest position, the distance between object table and objective front lens should be at least 5 cm. You will need that space for mounting all the accessories on top of the table. We use a Leica DMRE TCS SP2 microscope equipped with acousto-optical beam splitter (AOBS), three photomultiplier tubes for single-photon detection, two non-descanned detectors for multiphoton detection, and 20/0.7 (Fig. 19.2A) and 63/1.2 water immersion objectives corrected for use without coverglasses (Fig. 19.2O). As a general remark, always take care to use well-corrected objectives with numerical apertures as high as possible. This will enormously impact on your image quality and the amount of light needed for imaging. To directly couple the objective to the tissue of interest, we use artificial tear gel, which has similar optical properties as water, but does not easily drain off due to its high viscosity. Using such setup, the analysis of many different parameters in the living muscle can be executed. Limitations come from three main sources: probe availability, imaging speed, and spectral overlaps. But developments in probe generation and microscope technology continuously reduce the range of these limits. Due to the versatility of the approach, we can here only exemplify how multimodal imaging leads to more meaningful insights. We recently described that the localization of protein kinase A type I (PKA I) in close proximity to a subsynaptic cAMP microdomain is crucial for the proper turnover of acetylcholine receptors at that site (Ro¨der et al., 2010). To address this issue, we needed to correlate cAMP signals with synapse morphology and the presence of PKA I. Therefore, we transfected a FRET-based genetically encoded fluorescent cAMP sensor (RIa-EPAC), which also acts as an in vivo marker for PKA I localization. Ten days after transfection, we additionally injected the synaptic marker, a-bungarotoxin, coupled to the infrared dye, AlexaFluor 647. Then, using single-photon microscopy (Fig. 19.3A–D), we determined PKA I localization and synapse morphology. This showed the presence or absence of PKA RI in different synapses (Fig. 19.3A–C) and the size of these synapses in 3D (Fig. 19.3D). Subsequent multiphoton imaging (Fig. 19.3F–G) permitted to investigate the cAMP signaling in these fibers. Figure 19.3E shows one synapse
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Figure 19.3 Example for multifactorial in vivo imaging. Mouse tibialis anterior muscles were transfected with RIa-EPAC, a FRET-based sensor for cAMP that targets to a PKA RIa-specific microdomain in close proximity to neuromuscular junctions. Ten days after transfection, muscles were injected with fluorescently labeled a-bungarotoxin, a marker for acetylcholine receptors in neuromuscular junctions, and then imaged in vivo using confocal (A–E) and two-photon imaging (F, G). (A) Maximum z-projection of a 135 mm thick z-stack of optical slices taken at 3 mm interslice distance. The scale bar represents 50 mm. YFP-fluorescence signals of RIa-EPAC are shown in A2, a-bungarotoxin staining in A1. Individual optical sections of the boxed regions are shown in B and C. (B, C) Details of the optical section numbers 22 (B) and 28 (C) from the maximum z-projection depicted in A. The scale bar represents 50 mm. (B1 and C1) a-bungarotoxin staining signals. (B2 and C2) RIa-EPAC fluorescence signals. (D) Quantification of volumes of neuromuscular junctions a, b, and c. (E) Maximum z-projection of one synapse in single-photon mode. The scale bars represent 10 mm. Shown are the fluorescence signals of a-bungarotoxin staining (E1) and of YFP emitted from RIa-EPAC and (E2). (F1 and F2) YFP-fluorescence signals upon two-photon excitation at 820 nm of RIa-EPAC at different time points before and after injection of the cAMP agonist, forskolin (FRSK). (F3 and F4) Pseudocolored ratiometric images showing the ratio of CFP/YFP emission signals upon two-photon excitation at 820 nm. The scale bar beneath F4 indicates absolute ratio values. Note that only after application of FRSK a clear increase of the CFP/YFP ratio can be observed. (G) Graphical representation of the mean CFP/YFP ratio changes SEM (n ¼ 13 synapses) upon FRSK application.
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acquired in single-photon mode (synapse, Fig. 19.3E1 PKA I marker, Fig. 19.3E2), which was then analyzed by multiphoton imaging to determine its cAMP signals (Fig. 19.3F and G). Figure 19.3F1 and F2 show the yellow fluorescent protein (YFP) emission images upon multiphoton excitation at 820 nm at different time points before and after cAMP agonist application. Figure 19.3F3 and F4 depict the corresponding ratiometric images, which were created as described in Section 2.4. As already mentioned, combinations of this and other imaging modes allow to quantify a set of parameters in one and the same sample and therefore to correlate second messenger signals to their direct or indirect outputs. Due to the wide range of applications of multimodal imaging, we cannot discuss the requirements for every type of imaging. Therefore, in this chapter, we give only general remarks as to the acquisition of FRETbased second messenger signals. First, make sure that your additional dyes do not interfere with the FRET-fluorescence signals. We have made good experience with infrared stains, such as AlexaFluor 647 in combination with cyan fluorescent protein (CFP)–YFP FRET pairs (Fig. 19.3). The use of optical parameter oscillators (OPOs), which broaden the tunable multiphoton spectrum to about 1300 nm (current commercially available multiphoton lasers for microscopy stop at about 1000 nm wavelength), might extend the use of multimodal microscopy to additional dye combinations. Second, different types of FRET imaging can be pursued, but not all of them can be easily used for quantitative assessments. It is fair to say that FRET– fluorescence-lifetime imaging microscopy (FLIM) applications might be the most quantitative, but they are still pretty slow and, at least to our knowledge, they have not been described in living animals so far. Among the other available methods to detect FRET, we use dynamic ratiometric FRET imaging, which relies on increased FRET acceptor (e.g., YFP) emission at the expense of donor (e.g., CFP) emission upon donor excitation, when FRET efficiency is rising. Although it is not as quantitative as FRET–FLIM, simple division of data from simultaneously acquired image pairs (donor and acceptor emission) leads to quick and relatively movement-resistant insights into the dynamism of second messenger signals. In the case of CFP as a donor, multiphoton excitation at 820 nm has proven to bear very low crosstalk with YFP acceptors. Excitation spectra with samples expressing only one of the two FRET pair dyes permit identifying the wavelength where the amount of crosstalk is minimal. FRET image pairs should be taken at 12-bit image depth and sensors should be set to linearly respond to a wide range of incoming light. It is instrumental to work in the optimal gain range of your detectors and to keep these values constant over all your experiments. Also avoid using detector-offset values deviating strongly from zero, because this will lead to an increasingly nonlinear data acquisition. Finally, when using very different excitation and emission wavelengths, the inherent optical properties of lasers and lenses render a
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perfect alignment in x–y as well as in z very difficult. In other words, sequential use of short- and long-wavelength light might excite your sample at distinct planes. Additionally, the emission signals might undergo a differential movement along the optical path. These parameters should be checked for each new optical component, for example, using fluorescent beads. 2.3.1. Instrumentation and disposables needed 2.3.1.1. Instruments Upright microscope equipped with dual capabilities for measuring in singleand multiphoton mode Water immersion objectives for direct immersion with optimal correction for spherical and chromatic aberrations as well as high numerical aperture Custom-made support for mounting the animal onto the microscope object table Straight 9-cm surgical scissors Dumont #7 curved shanks forceps (11.5 cm) with and without serrations Narrow pattern curved forceps (12 cm) Hartman straight hemostat (10 cm) for fixing distal tendon to metal support Additional hardware (e.g., electrodes, force transducer) depends on the type of parameters to be analyzed 2.3.1.2. Solutions
Physiological solution to moisten the muscle during the preparation Anesthetics: Zoletil 100 (Virbac) and Rompun (2% xylazine, Bayer) Dye solutions in sterile physiological solution (if applicable) ViscOphtal (Winzer Pharma), artificial tear gel for objective immersion 2.3.1.3. Disposables
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2.4. Data extraction Data analysis again widely depends on which parameters need to be detected. For data analysis, we use ImageJ, a freeware, which was generated by the NIH (Wayne Rasband, NIH, Bethesda, USA) and now uses a broad base of customers, which participate in programming (http://rsb.info.nih.gov/ij/). We describe here the main features to analyze the dynamic FRET-based second messenger signals. First, subtract background from your images. In ImageJ, this can be done using the Process/Math/Subtract prompt. In most confocal or multiphoton applications, background is relatively low and stable and therefore a generic subtraction of a mean of background values measured outside the
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transfected area is mostly sufficient. Strong variations in background levels are often due to major changes in focus or due to alterations in tissue composition, such as upon influx of blood or lymph into the observed region. In such cases, a reliable background subtraction is difficult to achieve and data must be handled with extreme caution. If the origin of such deviations can be identified, try to eliminate or reduce it. After background subtraction, regions of interest need to be determined. Ideally, the signal of interest is so strongly above background that simply the entire signal above background can be used for data analysis. Alternatively, additional operations have to follow. We usually generate masks on the composites of the background-subtracted donor and acceptor images. For the composite, use the Add function under Process/ImageCalculator. Then, you may threshold pixels to be analyzed by using the Image/Adjust/ Threshold command. The Apply threshold command will lead to the creation of a mask, which, after setting minimum and maximum to 0 and 1, respectively, can be multiplied with the background-subtracted donor and acceptor images. The minimum–maximum setting can be achieved by dividing the mask images by 255. Multiplication with the donor and acceptor images can be done using the multiply function under Process/ImageCalculator. The resulting images can be used to create a ratiometric image file using the CalculatorPlus plugin. This allows dividing donor and acceptor images with an additional constant to avoid clipping of decimal places of ratio values (note that output values are always integers). For calculating the CFP/YFP ratio in the CalculatorPlus function Divide, i2 ¼ (i1/i2) k1 þ k2, we use donor and acceptor images as i1 and i2, respectively. This is good in sensors, where increase in a second messenger leads to reduced FRET, such as in EPAC sensors for cAMP. The inverse is the case with cameleon probes, where donor and acceptor images make i2 and i1, respectively. In all cases, a k1 value of 5000 avoids clipping of minima and maxima, and k2 is zero. Using such ratiometric images, the dynamics of second messenger signals can either be globally or locally determined. For global analyses, it is sufficient to read out the entire ratiometric images with a threshold higher than zero (in order to get rid of the background pixels). For local analyses, regions of interest might need to be adapted according to specific needs. For all these calculations good standard PCs are sufficient.
3. Adaptations to Tissues Other Than Skeletal Muscle 3.1. Isolated heart Single-photon confocal microscopy has extensively been used to study second messenger signaling in isolated cardiac cells. Use of a vast number of chemical- and protein-based reporters in single- and multimodal imaging experiments has allowed simultaneous interrogation of intracellular messengers including Ca2 þ, cAMP, or biophysical parameters such as
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membrane potential. The functional properties of cardiomyocytes, however, are heavily influenced by a number of factors including specific interactions with neighboring cardiomyocytes, fibroblasts, and the extracellular matrix, none of which are preserved in isolated cardiomyocytes in vitro. The ability to measure variations in intracellular second messengers at subcellular level within the intact tissue would thus provide key information to understand cellular physiology and pathophysiology of the heart. However, a relatively small number of studies have been conducted addressing second messenger imaging in rodent cardiac tissue (i.e., papillary muscles; Wier et al., 1997) or intact hearts, using either surface imaging (Minamikawa et al., 1997) or deep tissue imaging with multiphoton microscopy (Aistrup et al., 2006; Rubart et al., 2003). These models are especially relevant to infer the mechanism of cardiovascular diseases like heart failure, featuring extracellular matrix remodeling and fibroblast growth, or arrhythmias, for example, those resulting from defective cell to cell communication, whereby behavior of cardiomyocytes in their tissue environment may differ substantially from that of the isolated cell. Here, we illustrate an adaptation of the method described in detail above for skeletal muscle imaging to the use in the isolated, Langendorff-perfused mouse heart. We will focus on the protocol for multiphoton imaging with the Ca2 þ indicator fluo-4, although with slight variations, the same setup can be utilized with FRET-based sensors to detect intracellular levels of other second messengers. 3.1.1. Probe introduction into mouse hearts A number of different approaches can be used to obtain expression of fluorescent indicators in the mouse heart. These can be summarized as follows: transgenic expression, viral transduction either via systemic or local intramyocardial injection, and dye loading in the isolated heart. 3.1.2. Heart preparation Ten minutes after heparinization (heparin, 200 IU/kg, i.p.), the heart of a 2- to 3-month-old mouse is excised and the aorta cannulated with a blunt 18 G needle (Fig. 19.4A and B). Retrograde coronary perfusion of oxygenated Tyrode solution is achieved at room temperature, at a constant pressure of 70 mmHg, and carried on for 15–20 min to equilibrate. At this point, if dye loading is needed, a mixture of Fluo-4AM (10 mM)/ Pluronic F127 (10 mM) and sulfinpyrazone (SP, 10 mM) is added to the solution and perfused for 20 min, after which the heart is perfused with SP containing dye-free Tyrode solution for 30 min to allow de-esterification of the AM dye.
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Figure 19.4 Summary of the isolated heart Ca2 þ imaging experiment. (A) Schematic of isolated heart imaging. (B, C) Intact isolated mouse heart imaging involves three consecutive steps: (i) heart harvesting, aortic cannulation, and retrograde perfusion setup (B); (ii) dye loading by perfusion recirculation; and (iii) multiphoton imaging during electrical pacing (C). (D) Images recorded from subepicardial fibers of a fluo3AM-loaded mouse heart, during electrical pacing via the aortic cannula. Images were acquired at a rate of 8 frames/s during a systolic Ca2 þ transient and represented in intensity-weighed pseudocolor using a custom-made software routine operating in Matlab (Vimmaging, F. Mammano and C. Ciobutaru, Padova, Italy).
3.1.3. Heart imaging Tyrode solution perfusion is continued while the heart is mounted horizontally on a chamber made of an agarose cast (low melting point agarose, 4% in Tyrode; Fig. 19.4C), which is used to keep the heart in position under the microscope objective (Fig. 19.4A). To minimize movement, we use a combination of the contraction inhibitors butanedione monoxime (BDM, 10 mM) and blebbistatin (100 nM). A bipolar pulse generator (Ionoptix, USA) connected to the cannula and to a ground electrode embedded in the agarose cast is used to pace the heart. During imaging (Fig. 19.4C), a drop of tear gel is placed on the epicardial surface. When
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imaged with multiphoton microscopy, the first 8–10 subepicardial cardiomyocyte layers on the left ventricular wall appear oriented along the long axis of the heart and can be efficiently visualized (Fig. 19.4D). 3.1.4. Instrumentation and disposables needed 3.1.4.1. Instruments Upright microscope equipped with dual capabilities for measuring in singleand multiphoton mode Water immersion objectives for direct immersion with optimal correction for spherical and chromatic aberrations as well as high numerical aperture Straight 9-cm surgical scissors Dumont #7 curved shanks forceps (11.5 cm) Narrow pattern curved forceps (12 cm) Bipolar pulse isolator (i.e., MyoPacer Cell Stimulator, Ionoptix, USA) 22G Aortic Cannula for mouse heart (Cat No. SP3787, ADI Instruments, UK) 3.1.4.2. Solutions
Heart perfusion solution: HEPES-buffered Tyrode solution (HE-Tyrode), 100% O2 gassed Dye loading solution (if applicable): HE-Tyrode, Fluo-4AM (10 mM)/Pluronic F127 (10 mM), SP Dye washout solution: HE-Tyrode, SP Imaging solution: HE-Tyrode, BDM (10 mM), ()-Blebbistatin (100 nM, Tocris, MI, USA) Heparin, 200 IU/kg in PS Low melting point agarose ViscOphtal (Winzer Pharma), artificial tear gel for objective immersion
3.2. Brain imaging—A brief teaser Understanding signaling of neuronal circuits requires the monitoring of electric activities. Therefore, action potentials (APs) emitted by neurons are one of the most important functional parameters. APs have fast dynamics in the millisecond time scale and allow specific communication with other cells via spatio-temporal patterns. Because of their unparalleled temporal resolution, one would ideally utilize electrophysiological methods for intraand intercellular recordings within local networks and during behavior. However, the experimenter is faced with the challenge to apply them to behaving animals or to more than one cell at a time. An alternative approach and a fundamental breakthrough in general has been the use of multiphoton microscopy in combination with fluorescent probes sensitive to cellular processes, and appropriate for monitoring of synaptic activity (Dreosti and Lagnado, 2011; Grewe and Helmchen, 2009; Helmchen and Denk, 2005).
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Unfortunately, genetically encoded voltage-sensitive dyes, made by either fusing FPs to the Shaker Kþ channel (FlaSh), sodium channels, or built around voltage-sensitive phosphatases, still produce signals that are too small and slow to be used at the level of single cells. This limitation of low signalto-noise ratio needs to be overcome in the future by the development of new high-performance voltage sensors (Mutoh et al., 2011). However, in neurons, APs usually result in a transient increase in intracellular Ca2 þ concentrations. Thus, Ca2 þ imaging is currently the most promising approach for neuronal activity sampling in all cells within a region of hundreds of micrometers across, and at cellular resolution within local neuronal circuits (Grewe et al., 2010). Some studies have used bulk loading of membrane-permeant chemical Ca2 þ dyes, such as fluo-3, Oregon Green 488, or rhod-2. They can be introduced into thousands of cells through a small skull opening and pressure ejection (Garaschuk et al., 2006). This technique, termed multicell bolus loading, results in a rather uniform staining of neurons within the targeted area and also proved to be well suited for pharmacological studies in awake animals, for example, after local administration of drugs. However, since Ca2 þ transients form with a delay of a few milliseconds after the AP, and because Ca2 þ transients decay much slower than APs, Ca2 þ imaging has some limitation in the precise decoding of individual APs at high firing rates. In addition, bolus loading of chemical dyes lacks discrete indicator labeling of specific cell types. Recent studies have addressed these challenges. First, Grewe et al. (2010) greatly improved the temporal resolution of AP-evoked Ca2 þ transients for in vivo imaging in mouse neocortex down to 300 mm using random access scanning with acousto-optic deflectors (AOD). Notably, the AOD-based two-photon microscope allowed sampling rates of up to 500 Hz and determined spike times with near millisecond precision and only 5–15 ms confidence intervals. Although in vivo measurements were collected in two dimensions, this approach should be further extended using z-dimension scanning. Second, Drobizhev et al. (2011) characterized the two-photon absorption (2PA) spectra of 48 FPs, from enhanced BFP- and CFP-series, the “fruit” FPs to far-red variants, such as mRaspberry, mKate2, tandem dimer (td) Katushka2, or eqFP670. This approach will lead to the development of brighter and faster mutants for multiphoton applications. However, there are other factors of concern—such as photostability, photoswitching efficiency, and cellular expression rate. While some FP constructs like YC-cameleons under the control of constitutively active promoters proved to be critical (Nagai et al., 2004), similar indicators using troponin-C instead of calmodulin as a Ca2 þ sensing domain placed under the control of an inducible promoter were suitable to detect fast and robust Ca2 þ transients in neurons of transgenic mice (Hasan et al., 2004). However, the generation of transgenic mice is time consuming and often the overall success rate is moderate. A faster alternative is the use of lenti- or adenovirus-based systems for gene delivery to postmitotic cells. They offer several advantages,
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as stereotactic injection makes it easy to deliver recombinant probes locally and to control expression rate in a defined time window (Lutcke et al., 2010; Osten et al., 2006). Further, fluorescent labeling of a small population of neurons, in an otherwise unlabeled tissue, offers a high signal-to-noise ratio.
4. Concluding Remarks The monitoring of second messenger signaling in living animals and intact tissues is now an established method. In contrary to single cell-based assays, however, the modes of targeting the sensor into the tissue of interest and the challenges encountered during the imaging process greatly vary from tissue to tissue, and thus in each case, the methodological setup needs careful adaptation. This chapter elaborated on these aspects in three types of tissue, that is, skeletal muscle, heart muscle, and neuronal tissue to exemplify some of the most immediate practical challenges. We are aware that the concerns are likely to be different for other organs. In parallel with continuous refinement of the experimental procedures and with the generation of fluorescent probes more suited to multiphoton excitation, technical developments are rapidly leading to faster and more powerful microscopes. This combination is expected to enable in the near future increasingly better spatial and temporal resolution in the detection of biologically and pathologically relevant events in intact tissues and organs.
ACKNOWLEDGMENTS We apologize to all authors whose work could not be included due to space limitations. We are grateful to Anika Wagner for excellent technical assistance. The work of R. R. is supported by the Helmholtz Association, the Deutsche Forschungsgemeinschaft, and the Association Franc¸aise contre les Myopathies. Financial support from the Ministry of Science, Research and Art (BadenWu¨rttemberg—“Innovative Projekte”) to M. H. is gratefully acknowledged. M. M. is supported by the EU Seventh Framework Program FP7/2007-2013 (HEALTH-F2-2009-241526, EUTrigTreat).
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Dona, M., Sandri, M., Rossini, K., Dell’Aica, I., Podhorska-Okolow, M., and Carraro, U. (2003). Functional in vivo gene transfer into the myofibers of adult skeletal muscle. Biochem. Biophys. Res. Commun. 312, 1132–1138. Dreosti, E., and Lagnado, L. (2011). Optical reporters of synaptic activity in neural circuits. Exp. Physiol. 96, 4–12. Drobizhev, M., Makarov, N. S., Tillo, S. E., Hughes, T. E., and Rebane, A. (2011). Two-photon absorption properties of fluorescent proteins. Nat. Methods 8, 393–399. Garaschuk, O., Milos, R. I., and Konnerth, A. (2006). Targeted bulk-loading of fluorescent indicators for two-photon brain imaging in vivo. Nat. Protoc. 1, 380–386. Grewe, B. F., and Helmchen, F. (2009). Optical probing of neuronal ensemble activity. Curr. Opin. Neurobiol. 19, 520–529. Grewe, B. F., Langer, D., Kasper, H., Kampa, B. M., and Helmchen, F. (2010). High-speed in vivo calcium imaging reveals neuronal network activity with near-millisecond precision. Nat. Methods 7, 399–405. Hasan, M. T., Friedrich, R. W., Euler, T., Larkum, M. E., Giese, G., Both, M., Duebel, J., Waters, J., Bujard, H., Griesbeck, O., Tsien, R. Y., Nagai, T., et al. (2004). Functional fluorescent Ca2 þ indicator proteins in transgenic mice under TET control. PLoS Biol. 2, e163. Helmchen, F., and Denk, W. (2005). Deep tissue two-photon microscopy. Nat. Methods 2, 932–940. Lutcke, H., Murayama, M., Hahn, T., Margolis, D. J., Astori, S., Zum Alten Borgloh, S. M., Gobel, W., Yang, Y., Tang, W., Kugler, S., Sprengel, R., Nagai, T., et al. (2010). Optical recording of neuronal activity with a genetically-encoded calcium indicator in anesthetized and freely moving mice. Front. Neural Circuits 4, 9. Minamikawa, T., Cody, S. H., and Williams, D. A. (1997). In situ visualization of spontaneous calcium waves within perfused whole rat heart by confocal imaging. Am. J. Physiol. 272, H236–H243. Mutoh, H., Perron, A., Akemann, W., Iwamoto, Y., and Knopfel, T. (2011). Optogenetic monitoring of membrane potentials. Exp. Physiol. 96, 13–18. Nagai, T., Yamada, S., Tominaga, T., Ichikawa, M., and Miyawaki, A. (2004). Expanded dynamic range of fluorescent indicators for Ca(2þ) by circularly permuted yellow fluorescent proteins. Proc. Natl. Acad. Sci. USA 101, 10554–10559. Osten, P., Dittgen, T., and Licznerski, P. (2006). Lentivirus-based genetic manipulations in neurons in vivo. In “The Dynamic Synapse: Molecular Methods in Ionotropic Receptor Biology,” (J. T. Kittler and S. J. Moss eds.). CRC Press, Boca Raton, FL. Pozzan, T., Mongillo, M., and Rudolf, R. (2003). The Theodore Bucher lecture. Investigating signal transduction with genetically encoded fluorescent probes. Eur. J. Biochem. 270, 2343–2352. Ro¨der, I. V., Petersen, Y., Choi, K. R., Witzemann, V., Hammer, J. A., 3rd, and Rudolf, R. (2008). Role of Myosin Va in the plasticity of the vertebrate neuromuscular junction in vivo. PLoS One 3, e3871. Ro¨der, I. V., Lissandron, V., Martin, J., Petersen, Y., Di Benedetto, G., Zaccolo, M., and Rudolf, R. (2009). PKA microdomain organisation and cAMP handling in healthy and dystrophic muscle in vivo. Cell. Signal. 21, 819–826. Ro¨der, I. V., Choi, K. R., Reischl, M., Petersen, Y., Diefenbacher, M. E., Zaccolo, M., Pozzan, T., and Rudolf, R. (2010). Myosin Va cooperates with PKA RIalpha to mediate maintenance of the endplate in vivo. Proc. Natl. Acad. Sci. USA 107, 2031–2036. Romanello, V., Guadagnin, E., Gomes, L., Ro¨der, I., Sandri, C., Petersen, Y., Milan, G., Masiero, E., Del Piccolo, P., Foretz, M., Scorrano, L., Rudolf, R., et al. (2010). Mitochondrial fission and remodelling contributes to muscle atrophy. EMBO J. 29, 1774–1785.
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Rubart, M., Wang, E., Dunn, K. W., and Field, L. J. (2003). Two-photon molecular excitation imaging of Ca2 þ transients in Langendorff-perfused mouse hearts. Am. J. Physiol. Cell Physiol. 284, C1654–C1668. Rudolf, R., Mongillo, M., Magalhaes, P. J., and Pozzan, T. (2004). In vivo monitoring of Ca2 þ uptake into mitochondria of mouse skeletal muscle during contraction. J. Cell Biol. 166, 527–536. Rudolf, R., Magalhaes, P. J., and Pozzan, T. (2006). Direct in vivo monitoring of sarcoplasmic reticulum Ca2 þ and cytosolic cAMP dynamics in mouse skeletal muscle. J. Cell Biol. 173, 187–193. Tothova, J., Blaauw, B., Pallafacchina, G., Rudolf, R., Argentini, C., Reggiani, C., and Schiaffino, S. (2006). NFATc1 nucleocytoplasmic shuttling is controlled by nerve activity in skeletal muscle. J. Cell Sci. 119, 1604–1611. Valkova, C., Albrizio, M., Ro¨der, I. V., Schwake, M., Betto, R., Rudolf, R., and Kaether, C. (2011). Sorting receptor Rer1 controls surface expression of muscle acetylcholine receptors by ER retention of unassembled alpha-subunits. Proc. Natl. Acad. Sci. USA 108, 621–625. Whitaker, M. (2010). Genetically encoded probes for measurement of intracellular calcium. Methods Cell Biol. 99, 153–182. Wier, W. G., ter Keurs, H. E., Marban, E., Gao, W. D., and Balke, C. W. (1997). Ca2 þ ‘sparks’ and waves in intact ventricular muscle resolved by confocal imaging. Circ. Res. 81, 462–469. Zaccolo, M., and Pozzan, T. (2003). cAMP and Ca(2 þ) interplay: A matter of oscillation patterns. Trends Neurosci. 26, 53–55. Zaccolo, M., Magalhaes, P., and Pozzan, T. (2002). Compartmentalisation of cAMP and Ca (2 þ) signals. Curr. Opin. Cell Biol. 14, 160–166. Zhang, J., Campbell, R. E., Ting, A. Y., and Tsien, R. Y. (2002). Creating new fluorescent probes for cell biology. Nat. Rev. Mol. Cell Biol. 3, 906–918.